identiFicAtion - Invasive Species Research Institute

A GUIDE TO THE 

Identification 

and Control 

of Exotic 

Invasive Species 

IN ONTARIO’S HARDWOOD FORESTS 

Lisa M. Derickx and Pedro M. Antunes

A Guide to the 

Identification 

and Control 

of Exotic 

Invasive Species 

in ontArio’s hArdwood Forests 

Lisa M. Derickx and Pedro M. Antunes

Invasive Species Research Institute 

1520 Queen Street East, BT300 

Sault Ste. Marie, Ontario, Canada 

P6A 2G4 

© Invasive Species Research Institute 2013 

This publication is subject to copyright. No part of this publication may 

be reproduced, modified, or redistributed in any form or by any means, 

without the prior written permission of the authors. 

Algoma University 

ISBN 978-0-929100-21-0 

ISRI has been granted permission to reproduce the photographs and 

range maps in this book by their rightful copyright holders and through 

the Bugwood Network Image Archive. A full list of photography credits 

appears in section 8.0. 

Disclaimer: This publication contains recommendations for chemical 

control that are intended to serve only as a guide. It is the responsibility 

of the pesticide applicator to follow the directions on the pesticide label. 

Pesticide labels and registrations change periodically. If any information 

in these recommendations disagrees with the label, the recommendation 

must be disregarded. No endorsement is intended relating to pesticide 

use or a particular product. The authors and the Invasive Species 

Research Institute assume no liability resulting from the use of these 

recommendations. 

Design: Carmen Misasi Design 

Cover photo credits: English ivy leaves: James H. Miller, USDA Forest Service, Bugwood.org 

(left) & Chris Evans, Illinois Wildlife Action Plan, Bugwood.org (right); Japanese barberry, garlic 

mustard & Himalayan balsam: Lisa Derickx, Invasive Species Research Institute; Norway maple 

keys: Leslie J. Mehrhoff, University of Connecticut, Bugwood.org; tree-of-heaven samaras: Chuck 

Bargeron, University of Georgia, Bugwood.org; emerald ash borer adult & tunneling damage: 

David Cappaert, Michigan State University, Bugwood.org; emerald ash borer feeding galleries: 

Joseph O’Brien, USDA Forest Service, Bugwood.org; Asian long-horned beetle: Melody Keena, 

USDA Forest Service, Bugwood.org; beech bark disease: Linda Haugen, USDA Forest Service, 

Bugwood.org; gypsy moth larva: Jeffrey Fengler, Connecticut Agricultural Experiment Station 

Archive, Connecticut Agricultural Experiment Station, Bugwood.org; Dutch elm disease: Fabio 

Stergulc, Università di Udine, Bugwood.org; crown dieback and epicormic shoots: Daniel Herms, 

The Ohio State University, Bugwood.org 

Back cover photo credits: Emerald ash borer feeding on leaf: Jared Spokowsky, New York State 

Department of Agriculture and Markets, Bugwood.org; Asian long-horned beetle exit hole: Steven 

Katovich, USDA Forest Service, Bugwood.org; Asian long-horned beetle feeding damage: Dean 

Morewood, Health Canada, Bugwood.org; elm bark beetle larval galleries: James Solomon, USDA 

Forest Service, Bugwood.org; butternut canker: Joseph O’Brien, USDA Forest Service, Bugwood. 

org; dog-strangling vine seedpod: Leslie J. Mehrhoff, University of Connecticut, Bugwood.org; 

common buckthorn, Japanese knotweed, periwinkle, dog-strangling vine flowers and exotic 

bush honeysuckle: Lisa Derickx, Invasive Species Research Institute.

This book is dedicated to Errol Caldwell for everything he does 

for Northern Ontario’s social and economic development. The 

creation of an Invasive Species Research Institute at Algoma 

University in Sault Ste. Marie and this guidebook were only 

possible due to his foresight.

[ iv ] 

Acknowledgments 

Funding for this book was provided by the Invasive Species 

Centre (ISC) to the Invasive Species Research Institute (ISRI) 

at Algoma University. As a not-for-profit organization, part of 

ISRI’s mission is to ensure that our research findings reach 

as many people as possible. In the case of this publication, 

the external funding received was essential to produce this 

guidebook and to print and distribute it at the lowest possible 

cost (i.e., only that of printing). 

We are thankful to Dr. Michael Irvine, Vegetation Management 

Specialist from the Ontario Ministry of Natural Resources 

(OMNR), for providing constructive criticism throughout the 

development of this guide. His consultation in the early stages 

of writing and his thorough review of the manuscript have 

lent to the significance and value of this publication. Thanks 

to Professor John Klironomos from the University of British 

Columbia for writing the foreword; to Dr. Richard Wilson, Forest 

Program Pathologist (OMNR), for his expert advice and helpful 

suggestions regarding forest pathogens; to Dr. W.D. McIlveen, 

Terrestrial Biologist, and Susan Meades, Director of the Northern 

Ontario Plants Database, for their advice and review of plant 

species taxonomy; to the ISRI staff, Laura Sanderson and Kim 

Mihell, for proof reading the text and assistance with editing. 

We are grateful to Jeff and JoAnn St. Pierre from North Country 

Photography and James Smedley from James Smedley Outdoors 

for assistance in photographing all aspects relating to maple 

syrup production. A special thanks is extended to the many 

other people who provided permission to use the high quality 

photographs found in this guide. A full list of photo credits 

follows to acknowledge these contributions. We are also 

thankful to our designer, Carmen Misasi of Carmen Misasi 

Design, for all of the time and effort he put into working with 

us to create this publication. 

We are thankful to Calvin Gilbertson (Gilbertson’s Maple 

Products) and David Thompson (Thompson’s Maple Products) 

for permission to photograph their maple syrup operations 

on St. Joseph Island and for sharing their expert knowledge of 

maple syrup production in Ontario. Also, thanks to the Ontario 

Maple Syrup Producers Association (OMSPA) and all of their 

members for a wonderful and educational experience at the 

summer tour and meeting in Belleville, 2011.

Thanks to Lindsay Burtenshaw at the Royal Botanical Gardens 

and Bruce Cullen from the Toronto Zoo for their information 

on managing invasive plants, for tours of the gardens and 

zoo, and for permission to take photographs for use in the 

guide. Thanks to Freyja Forsyth of Credit Valley Conservation 

(CVC), Hayley Anderson of the Ontario Invasive Plant Council 

(OIPC) and Fraser Smith of the Ontario Federation of Anglers 

and Hunters (OFAH) Invading Species Awareness Program, 

for providing us with hands-on experience in invasive plant 

management at their various volunteer events. 

Finally, thanks to our family and friends for their patience and 

encouragement over the course of producing this guide. 

[ v ]

[ vi ] 

About the Authors 

Lisa M. Derickx, B.Sc., is a Research Associate at the Invasive 

Species Research Institute (ISRI) in Sault Ste. Marie, Ontario. 

She has a B.Sc. Honours Degree in Environmental Science 

from Carleton University and a diploma in Fish and Wildlife 

Conservation from Sault College of Applied Arts and Technology. 

Lisa has obtained funding from the Ontario Trillium Foundation 

to develop a citizen scientist approach to identify and map 

terrestrial invasive plants. Her interests lie in plant and wildlife 

identification and environmental conservation. 

Pedro M. Antunes, B.Sc. & Ph.D, is a Research Chair in Invasive 

Species Biology (funded by the Ontario Ministry of Natural 

Resources) and Associate Professor (Department of Biology, 

Algoma University) since 2010. Currently, he is also the 

Research Director of the Invasive Species Research Institute at 

Algoma University and Chair of the North American Invasive 

Species Network. He began his academic studies in Biology at 

the University of Évora, Portugal (1999). He then undertook his 

doctoral research in Soil Science at the University of Guelph 

(2005) followed by post-doctoral research in Soil Microbial and 

Plant Ecology (2005-07). In 2008, he moved to Berlin, Germany, 

to assume a Research Assistant Professor position in Ecology 

at the Freie Universität. He is broadly interested in science and 

environmental conservation. His research in ecology focuses 

on the roles that soil microorganisms play in controlling plant 

productivity and community structure.

Foreword 

As humans, we love to place things into groups. It is how we 

organize the various objects that we come into contact with, 

it is the filing system that we have for our thoughts. It is how 

we make sense of the world. When taking a walk in nature, we 

may not know how to identify every organisms that we see, but 

we can typically divide them into broad groups: Plant, animal, 

fungus - and from there (and often with the help of a field 

guide) we can get more specific, a particular Phylum, and then 

all the way down to a species. 

The species binomial (or latin name) is the gateway to all 

information that is known for that organism - its life history, its 

morphological and behavioural characteristics, its geographic 

range, and whether it is native or exotic. Interestingly, a large 

proportion of the species that we encounter in most habitats are 

exotic. They originated elsewhere and immigrated to our local 

ecosystems. Where did these species come from? Well, the vast 

majority of exotic organisms that are found in Ontario have 

origins in Eurasia, from locations with comparable climate. 

If we want to learn more about these species, then a visit to 

their home of origin may provide significant insight into their 

biology and ecology. 

Why worry about exotic organisms? Is this a form of 

xenophobia? One major reason for concern is that they lead to 

the homogenization of our natural ecosystems. Everywhere we 

go, we see the same species, diminishing the uniqueness of local 

ecosystems. As humans, this has a strong psychological effect 

on us. It is unnerving, similar to when we travel to a distant 

location overseas and find the same fast food restaurants that 

we just left behind. We don’t yet understand the ecological 

consequences of such homogenization. However we do know 

that many exotic organisms, those that are invasive, can have 

great ecological and economic consequences. 

Being exotic is one thing. Being invasive is another. It is 

important not to confuse the two. All invasive organisms are 

exotic but not all exotics are invasive. How do we define an 

invasive species? There are two main categories. The first are 

those species that grow, reproduce, and spread at unusually 

high rates. Such species can be found at very high densities in 

local habitats, often in widespread monoculture. The second 

are those species that have significant local impact. They can 

negatively affect the growth and reproduction of local native 

species, or may alter the functioning (productivity, stability) of 

local ecosystems. Some of the most problematic and worrisome 

invasive species can do both, spread rapidly and harm native 

systems. 

[ vii ]

[ viii ] 

Scientists are currently placing a lot of effort in the study of 

invasive species. There are many unanswered questions, such 

as: what factors cause certain species to become invasive? Why 

are certain ecosystems more invasible than others? What are 

effective ways to control invasive species? Such questions are not 

just academic. We need to know the answers to these questions, 

so that we can properly manage our landscapes. In this context, 

any new information on invasive species, particularly the most 

serious of them, needs to be communicated to other scientists 

and to the public. So it should be clear just how important this 

book is. A guide that focuses on invasive species in Ontario - 

one that is comprehensive and science-based, yet accessible to 

any audience. 

Dr. John Klironomos 

University of British Columbia

Dedication...............................................................................................................................................................................................iii 

Acknowledgments..................................................................................................................................................................................iv 

About.the.Authors..................................................................................................................................................................................vi 

Foreword.............................................................................................................................................................................................................vii 

1.0. Introduction 

section one 

. 1.1. An.Introduction.to.Exotic.Invasive.Species.of.Hardwood.Forests........................................................................2 

. 1.2. How.to.Use.this.Book...........................................................................................................................................................4 

2.0. Invasive.Species.Management:.An.Overview 

. 2.1.. Prevention.Strategies...........................................................................................................................................................8 

. 2.2. Early.Detection.and.Rapid.Response............................................................................................................................11 

. 2.3. Management.and.Control.Options...............................................................................................................................13 

. . 2.3.1. Physical.Control......................................................................................................................................................13 

. . 2.3.2. Chemical.Control....................................................................................................................................................16 

3.0. A.Tribute.to.Hardwood.Forests.in.Ontario 

. 3.1. Timber.Harvest.................................................................................................................................................................... 20 

. . 3.1.1. Ontario’s.Forests.................................................................................................................................................... 20 

. . 3.1.2. History,.Economy.and.Culture.of.Timber.Harvest...................................................................................... 22 

. 3.2. Maple.Syrup.Production.................................................................................................................................................. 23 

. . 3.2.1. History....................................................................................................................................................................... 23 

. . 3.2.2. Economic.and.Cultural.Significance............................................................................................................... 28 

4.0. Woodlot.Management 

Table of Contents 

. 4.1. Management.for.Timber.Harvest..................................................................................................................................31 

. 4.2. Sugar.Bush.Management................................................................................................................................................ 34 

[ ix ]

[ x ] 

5.0.Invasive.Species.Accounts 

section two 

. 5.1. Exotic.Plants......................................................................................................................................................................... 38 

. . 5.1.1. Norway.Maple.(Acer platanoides)..................................................................................................................... 39 

. . 5.1.2. Tree-of-heaven.(Ailanthus altissima)............................................................................................................... 53 

. . 5.1.3. Garlic.Mustard.(Alliaria petiolata)..................................................................................................................... 69 

. . 5.1.4. Barberry.(Berberis thunbergii.&.B. vulgaris).................................................................................................... 85 

. . 5.1.5. Japanese.Knotweed.(Fallopia japonica)......................................................................................................... 97 

. . 5.1.6. English.Ivy.(Hedera helix)...................................................................................................................................109 

. . 5.1.7. Himalayan.Balsam.(Impatiens glandulifera)................................................................................................ 121 

. . 5.1.8. Common.Buckthorn.(Rhamnus cathartica)................................................................................................. 131 

. . 5.1.9. Periwinkle.(Vinca minor).................................................................................................................................... 143 

. . 5.1.10. Dog-strangling.Vine.(Vincetoxicum rossicum.&.V. nigrum)..................................................................... 155 

. . 5.1.11. Goutweed.(Aegopodium podagraria)........................................................................................................... 171 

. . 5.1.12. Oriental.Bittersweet.(Celastrus orbiculatus)................................................................................................ 173 

. . 5.1.13. Exotic.Bush.Honeysuckle.(Lonicera.spp.)..................................................................................................... 175 

. . 5.1.14. Kudzu.(Pueraria montana.var..lobata)........................................................................................................... 177 

. 5.2. Exotic.Insects.&.Disease.................................................................................................................................................180 

. . 5.2.1. Emerald.Ash.Borer.(Agrilus planipennis)....................................................................................................... 181 

. . 5.2.2. Asian.Long-horned.Beetle.(Anoplophora glabripennis).......................................................................... 191 

. . 5.2.3. Chestnut.Blight.(Cryphonectria parasitica)..................................................................................................203 

. . 5.2.4. Beech.Bark.Disease.Complex.(Cryptococcus fagisuga.&.Neonectria.spp.)......................................... 213 

. . 5.2.5. Gypsy.Moth.(Lymantria dispar)........................................................................................................................223 

. . 5.2.6. Dutch.Elm.Disease.(Ophiostoma.spp.).......................................................................................................... 231 

. . 5.2.7. Elm.Bark.Beetles.(Scolytus multistriatus.&.S. schevyrewi)........................................................................ 241 

. . 5.2.8. Butternut.Canker.(Ophiognomonia clavigignenti-juglandacearum)...................................................249 

. . 5.2.9. Dogwood.Anthracnose.(Discula destructiva).............................................................................................257 

. . 5.2.10. Thousand.Cankers.Disease.(Geosmithia morbida)....................................................................................259 

. . 5.2.11. Pear.Thrips.(Taeniothrips inconsequens)....................................................................................................... 261 

6.0. Appendices.................................................................................................................................................................................263 

7.0. References...................................................................................................................................................................................267 

8.0. Photography.Credits................................................................................................................................................................281 

9.0. Acronyms. ...................................................................................................................................................................................283

1.0 Introduction 

2.0 Invasive Species 

Management: An 

Overview 

3.0 A Tribute to Hardwood 

Forests of Ontario 

4.0 Woodlot Management

[ 2 ] 

A GUIDE TO THE IDENTIFICATION AND CONTROL OF EXOTIC INVASIVE SPECIES IN ONTARIO’S HARDWOOD FORESTS 

1.0 

introduction 

1.1 

An introduction to exotic invasive 

species of hardwood Forests 

Exotic invasive species are those not native to the habitat 

in which they are causing harm to either the environment, 

economy or society (Environment Canada, 2004). Invasive 

species are increasingly prevalent in Ontario’s hardwood 

forests. Forestry represents an important part of Ontario’s 

economy, at an estimated $14 billion in 2008 (MNDM, 2011). 

Ontario’s hardwood forests also provide high value non-timber 

forest products such as the iconic maple syrup (Mohammed, 

1999) (Fig. 1). They provide us with ecological goods and 

Figure 1: A non-timber forest product (sap for ‘maple syrup’) 2.

INTRODUCTION 1.0 

services, such as clean air and wildlife habitat, provide areas 

for recreational enjoyment, contribute to the natural aesthetics 

of Ontario and sustain a resource-based tourism industry. For 

all these reasons, proper management of invasive species is a 

vital step to sustaining our existing hardwood forests (Daily et 

al. 1997). 

Invasive species can alter forest integrity through rapid 

population expansion. This may affect biodiversity by causing 

shifts in species abundances and in some cases can lead to local 

extinctions (Wyckoff & Webb, 1996). Studies have shown that 

exotic plant invasions can alter soil physicochemical properties 

and affect nutrient cycling (Leicht-Young et al. 2009; Kourtev 

et al. 1998). Moreover, exotic insects and pathogens (i.e., 

microorganisms that cause disease in their hosts) can severely 

damage and cause large-scale mortality of indigenous trees and 

shrubs (Allen & Humble, 2002). Consequently, invasive species 

can result in reductions to timber value and the quality and 

quantity of other forest-derived products such as maple syrup 

(Kota et al. 2007). 

The majority of exotic plants were introduced to North America 

intentionally for agriculture and horticultural purposes and 

were thus prized for their fast-growing, sun-loving habits. 

Since forest understories are generally subject to low light 

levels, they tend to be inhospitable to many of these introduced 

horticultural varieties. However, some shade-tolerant exotic 

species have been introduced and, not surprisingly, many are 

spreading in forest environments (Martin et al. 2008). 

In general, invasive forest plants have relatively long lagtimes 

(i.e., the time it takes before a species’ population grows 

exponentially and becomes invasive in the new introduced 

environment), which may give a false sense of security when 

assessing the risk of invasion (Crooks, 2005). Some of the 

invasive plants described in this book are still available for sale 

in garden centres across Ontario. Only recently has evidence 

arisen concerning the detrimental effects of invasive vines 

such as periwinkle (Vinca minor; Darcy & Burkart, 2002) and 

English ivy (Hedera helix; Dlugosch, 2005). Other species, such 

as common barberry (Berberis vulgaris), have been banned for 

sale due to reported detrimental effects to agriculture; yet its 

close relative, Japanese barberry (B. thunbergii), is still sold and 

used as a common garden plant (Fig. 2) because it is not known 

to cause any ill effects to agricultural crops (CFIA, 2008). More 

research is required to determine the factors responsible 

for different lag-times in exotic plant species. However, the 

exotic status should be sufficient for the inclusion of these 

species in monitoring programs (Simberloff, 2011). For highly 

invasive species, research on their ecological effects as well 

as preventative measures need to be considered to minimize 

negative effects to hardwood stands in Ontario. 

[ 3 ]

[ 4 ] 


Figure 2: Two varieties of a common garden shrub, Japanese barberry 6 . 

1.2 

how to use this Book 

Considering the increasing number of invasive species establishing 

in Ontario’s forests, there is a clear need for management 

guidelines focusing on those species that have the largest 

potential to be detrimental to hardwood stands. In addition, 

challenging economic times are leading to increasing awareness 

that public involvement in environmental issues is crucial 

to overcome these challenges. This guide, based on evidence 

from the peer-reviewed scientific literature, focuses on invasive 

plants, insects and pathogens that exhibit ecological, economic 

and social impacts that are detrimental to Ontario’s hardwood 

forests. It is intended to serve as an educational resource and a 

field guide to aid woodlot owners in invasive species management.


There are two sections in this guide. The first section describes 

the risks associated with invasive species in hardwood forests. 

These forests are valuable to woodlot owners in Ontario who 

rely on them for timber and high-valued products such as 

maple syrup. It introduces managers to the basic principles 

associated with different management strategies and outlines 

various methods of invasive species control. 

Species invasions are dynamic and each case is unique. 

Management recommendations in this guide may not be the 

most appropriate for everyone. Forest managers and woodlot 

owners should gauge their level of expertise, commitment and 

finances when considering control options. It is also important 

to understand that these recommendations and management 

strategies may change as ongoing research provides new 

insights into invasive species management. We point out gaps 

in the literature throughout the guide and recommend that the 

reader keeps informed on highly invasive species and up-todate 

with current management practices. 

Individual invasive species descriptions make up the second 

section of this guide. These include invasive plants, insects 

and pathogens that have been identified as priorities for 

management in Ontario’s hardwood stands (Tables 1 & 2). They 

have been selected as priorities for management based on the 

following risk categories: 

Economic risks: 

• Species that can directly affect commercial hardwoods by 

significantly reducing growth, causing widespread mortality 

and/or suppressing hardwood regeneration; 

• Species that can alter the quality of timber or the quantity of 

maple syrup produced; 

• Species whose environmental effects can result in the 

reduction of property value; 

• Species that can cause losses resulting from movement 

restrictions related to timber products. 

Environmental risks: 

• Species that cause the loss of biodiversity; 

• Species that reduce ecosystem goods and services; 

• Species that can harm a species at risk. 

Social impacts: 

• Species that can interfere with traditional lifestyles; 

• Species that can reduce aesthetic values of a hardwood stand; 

• Species that affect the recreational enjoyment of a woodlot; 

• Species that can impact human health. 

[ 5 ]

[ 6 ] 


Table 1: Invasive plants considered a high priority for management in hardwood stands and their 

associated risks (see Appendix 1 for referenced priority rating). 

Norway maple 

(Acer platanoides) 

Tree-of-heaven 

(Ailanthus altissima) 

Garlic mustard 

(Alliaria petiolata) 

Barberry 

(Berberis spp.) 

English ivy 

(Hedera helix) 

Himalayan balsam 

(Impatiens glandulifera) 

Japanese knotweed 

(Fallopia japonica) 

Common buckthorn 

(Rhamnus cathartica) 

Periwinkle 

(Vinca minor) 

Dog-strangling vine 

(Vincetoxicum spp.) 

Cause reduced 

hardwood growth 

Economic Risks Environmental Risks Social Risks 

Cause hardwood 

mortality 

Suppress hardwood 

regeneration 

Cause the loss of 

biodiversity 

Affect ecosystem 

function 

Harm species-atrisk 

Interfere with a 

traditional lifestyle 

Reduce aesthetic 

values of the forest 

The following information will be provided for invasive species 

considered a high priority in hardwood stands: 

Identification – A tool to help managers accurately identify 

invasive species present in their woodlots. Signs and symptoms 

are provided to help identify invasive insects and pathogens. 

Similar species – Visual aids and dichotomous keys are included 

to help distinguish similar species. 

Biology – Understanding the biology and taxonomy of an 

invasive species provides important clues for its management. 

Taxonomic hierarchy, origin and distribution, habitat, type 

of reproduction, lifecycle and host species are included for 

applicable species. 

Success mechanisms – Understanding traits likely to play an 

important role in enabling species invasibility can help managers 

Affect recreational 

enjoyment 

Impact human 

health

table 2: Invasive insects and pathogens considered a high priority for management in hardwood stands 

and their associated risks (see Appendix 1 for referenced priority ratings). 

Emerald ash borer 

(Agrilus planipennis) 

Asian long-horned beetle 

(Anoplophora glabripennis) 

Chestnut blight 

(Cryphonectria parasitica) 

Gypsy moth 

(Lymantria dispar) 

Beech bark disease 

(Neonectria faginata) 

Dutch elm disease 

(Ophiostoma spp.) 

Elm bark beetles 

(Scolytus spp.) 

Butternut canker 

(Ophiognomonia clavigignentijuglandacearum) 

Cause reduced 

hardwood growth 


Economic Risks Environmental Risks Social Risks 

Cause hardwood 

mortality 

Suppress hardwood 

regeneration 

Cause the loss of 

biodiversity 

Affect ecosystem 

function 

Harm species-at-risk 

Interfere with a 

traditional lifestyle 

Reduce aesthetic 

values of the forest 

with the decision making process to select appropriate invasive 

species control strategies. 

Ecological impacts – Understanding the ecological effects of 

invasive species is the basis for why they need to be managed. 

Vectors and pathways – Understanding how and where an 

invasive species can reach and establish in a woodlot can 

greatly improve prevention and early detection techniques. 

Management practices 

• Prevention strategies – These strategies are the most efficient 

and cost effective methods of invasive species management. 

These strategies outline various ways whereby managers can 

limit invasive species introductions to woodlots. 

• Early detection techniques – These techniques help maximize 

the capacity to detect invasive species before they spread in 

the woodlot. 

• Control options – Outlines the most cost effective and environmentally 

sustainable management strategies for priority 

species. 

Affect recreational 

enjoyment 

Impact human 

health 

[ 7 ]

[ 8 ] 


2.0 

invAsive sPecies MAnAGeMent: 

An overview 

2.1 

Prevention strategies 

Prevention is the most efficient and effective approach to 

manage exotic invasive species. There are several things a 

manager can do to prevent the introduction of invasive species 

into a woodlot, regardless of whether they are plants, insects 

or pathogens. Creating a prevention plan that involves all those 

responsible for woodlot activities can go a long way to reducing 

the risk of invasion and the amount of time, effort and cost 

needed for invasive species management (Clark, 2003). 

Prevention strAteGies 

For invAsive PlAnts 

• Promote a healthy forest with diverse communities of native 

species. An undisturbed forest understory with diverse 

plant communities may be more resistant to exotic species 

introductions by imposing a high level of competition on 

newcomers (Clark, 2003). Remember to monitor both the 

canopy trees and the understory vegetation. 

• Minimizing the agents of invasive species dispersal (i.e., 

vectors) will decrease the likelihood of introduction. Humans 

are a main vector of seed dispersal, so limiting access to the 

woodlot may be beneficial (Wilkins, 2000). However, this may 

not be possible in hardwood stands used for maple syrup 

production where access is required during the sugar season.

INVASIVE SPECIES MANAGEMENT: AN OVERVIEW 2.0 

In this case, access should be restricted to existing trails and 

roads. Ensuring that shoes and tires are cleaned before entry 

could save a manager time and money (NCC, 2007). 

• When planning for timber harvest, ensure that prevention 

measures are considered in the work plan. Survey the area to 

locate any existing invasions. Where possible, remove these 

species several years before harvesting. If eradication is not 

feasible, use an appropriate method of control. This will 

ensure that invasive plants are managed before activities that 

could contribute to further spread take place (Clark, 2003). 

• Soil disturbance increases the forest’s susceptibility to plant 

invasions. Try to minimize activities such as road or trail 

construction. As a mitigation strategy, plant native species 

where soil disturbance is unavoidable. This may help to deter 

establishment of invasive plants by promoting competition 

(Clark, 2003). 

• Any equipment that must be brought into the woodlot such 

as tractors, skidders and all-terrain vehicles should be 

thoroughly cleaned. All mud and plant materials should be 

washed off in a designated area before entering the woodlot. 

These designated areas should be monitored frequently for 

invasive plants (USDA, 2001). 

• Be careful when bringing any materials such as sand, gravel 

or soil into the woodlot. If possible, managers should take 

the initiative to visit the seller’s warehouse beforehand 

to inspect materials for seedlings of invasive species. Ask 

companies whether they have implemented measures to 

prevent the transportation of invasive plants (USDA, 2001). 

• Using the woodlot for recreational purposes such as hiking 

can also be a potential vector for seed dispersal. Shoes should 

be washed before entering and upon leaving the woodlot at a 

designated area. This area must also be frequently monitored 

for germinating seeds (NCC, 2007). 

• Whenever possible keep pets on a leash because seeds can 

easily get caught in fur. This is especially important during 

the late summer and fall when seeds are ready to disperse 

(Clark, 2003). 

• Avoid walking or driving through invaded areas. Traveling 

through these areas can spread seeds to un-invaded areas of 

the woodlot. Consider using signs such as flags to prevent 

others from walking through the invaded area (Clark, 2003). 

• Continue to be conscientious outside the boundaries of the 

woodlot. Never plant an exotic invasive species in the garden 

as seeds can easily be carried into your woodlot (Miller et al. 

[ 9 ]

[ 10 ] 


2010). Ask local greenhouses about the possibility of buying 

native plants for the home garden (Havinga, 2000). 

• Volunteer to help control invasive plants on neighbouring 

properties. It will help to decrease the likelihood of seed 

dispersal into your woodlot and will promote cooperation 

should a population establish on your own property (Peterson, 

2007). 

Prevention strAteGies For 

invAsive insects And PAthoGens 

• Increase species diversity in the woodlot. Many invasive 

insects and pathogens have a single host genus or species. 

Woodlots that are composed of one or two species are more 

likely to become infested and suffer greater negative effects 

when invasions do occur (OMNR, 2011). 

• Don’t move firewood. Many areas in Ontario have been put 

under various quarantines that restrict the movement of 

firewood and other wood products because they can harbour 

insects and pathogens. Check with the CFIA for quarantines 

in effect in your area (CFIA, 2012a). 

• Maintain a healthy woodlot. Perform thinning activities when 

required to help decrease both interspecifc and intraspecific 

competition around high value trees, thereby promoting 

vigorous tree growth. Pruning high-value trees to remove 

dead or damaged branches may also decrease pathogen 

infections (Ostry et al. 1996). 

• Avoid damaging trees during woodlot management activities. 

Wounds in the bark increase the chances of pathogen 

infection or insect establishment (Ostry et al. 1996). 

• Inspect and clean any vehicles, camping equipment, boats or 

other objects that may harbour invasive insects, especially 

when moving through quarantine zones (CFIA, 2011a).


2.2 

early detection and rapid response 

Although a prevention plan is considered an effective 

management approach, these are not always foolproof and 

it is best to have a backup plan in place (Clout & Williams, 

2009). Early detection is the identification of newly established 

invasive species at the initial stage of invasion. Invasive 

species may be eradicated and are clearly easier to control in 

these early stages. The larger a population is the greater the 

amount of labour and money required for its management 

(O’Neil et al. 2007). As such, management actions should 

occur promptly after detection to save money and minimize 

damage to the woodlot (NCC, 2007). 

Monitoring is the key to early detection. A monitoring 

program is essential and can be carried out by managers, 

friends, family members or volunteers. It involves actively 

searching for invasive species in the woodlot. Be aware of 

invasive species present on adjoining properties because they 

can spread quickly (NCC, 2007). Once an invasive species is 

detected, implementing actions to immediately address the 

problem is highly recommended. Having a plan that outlines 

the management options for highly invasive and thus priority 

species will allow for quick and efficient control of new 

invasions (Grice, 2009). 

eArly detection techniques 

For invAsive PlAnts 

• Early detection is important to prevent the establishment 

of an invasive plant species population and the consequent 

creation of a large seed bank. Remember that it is much 

easier to control a few individuals than a large population 

(Drayton & Primack, 1999). 

• Learn how to properly identify priority invasive plant 

species. Become familiar with those species that are already 

present in your region (Strobl & Bland, 2000). Educate family, 

friends and employees on how to properly identify priority 

invasive species (Siegal & Donaldson, 2003). 

[ 11 ]

[ 12 ] 


• Create a species inventory of the woodlot. Many managers 

may already have tree species inventories as part of their 

woodlot management plan. These inventories should also 

include understory species. Create a basic map of the different 

species present in the woodlot. With that, if any new species 

appears, it can be more readily detected. These measures will 

decrease the likelihood of missing new invasions (Miller et 

al. 2010). 

• Organize a monitoring program for invasive species. 

Monitoring activities should occur as frequently as possible 

and at least once during the spring, summer and fall (Miller 

et al. 2010). Pay particular attention to high risk areas such 

as forest edges, trails, roadways, flood water paths and 

disturbed areas. These are the pathways through which 

seeds can more easily disperse (Drayton & Primack, 1999). 

eArly detection techniques For 

invAsive insects And PAthoGens 

• Visual surveys and monitoring activities are important in 

the detection of invasive insects and pathogens in the early 

stages of invasion. Knowing what to look for is crucial. 

Invasive insects and pathogens usually leave visible signs of 

their presence in host trees (Cappaert et al. 2005). 

• Check for signs of insect infestation or disease symptoms on 

host trees. Do not limit monitoring to the more valuable trees 

because many insects and pathogens will attack weakened or 

dying trees first (Seybold et al. 2008). 

• Monitor host trees at various heights within the forest 

structure. Do not limit searches to what can be seen at eye 

level. Remember to pay attention to both the main stem and 

branches within the canopy (Turgeon et al. 2010). 

• Look for multiple signs of invasion to help properly identify 

the causative agent. The signs and symptoms of harmful 

invasive species can easily be mistaken for those of less 

harmful native species and vice-versa. Symptoms caused by 

various environmental stresses may also be mistaken for 

invasions (Turgeon et al. 2010).


2.3 

Management and control options 

Management and control options for invasive species that 

affect hardwood stands should be based on maintaining a 

healthy forest. A healthy forest will provide the economic 

and aesthetic benefits desired by managers (Havinga, 2000). 

Effective management involves planning a strategy and having 

the ability to provide a certain level of commitment, effort and 

money (Clout & Williams, 2009; Holcombe & Stohlgren, 2009). 

The following sections outline the three methods of invasive 

species control, including physical, chemical and biological 

control. Physical control consists of a person mechanically 

removing or destroying the invasive species with the use of 

hands or tools. Chemical control consists of using pesticides. 

Biological control consists of releasing living organisms that 

feed upon, parasitize or infect the unwanted species. When using 

chemical control an integrated approach to invasive species 

management should always be used. Integrated management 

requires thorough knowledge of the invasive species. All 

options including both mechanical and chemical control should 

be considered, taking into consideration all of the potential 

environmental and social risks, before a management strategy 

is implemented. Forest managers and woodlot owners should 

choose methods that are well aligned with their objectives and 

goals for the present and future use of the woodlot. 

2.3.1 

Physical control 

There are several ways of physically controlling invasive plants 

in a hardwood stand. They include hand-pulling, excavation, 

flower-head removal, cutting, mulching, solarization and using 

a directed flame. Hand-pulling consists of physically pulling 

whole plants, including the root system, out of the ground 

(Fig. 3). This is easiest when the soil is wet. Hand-pulling can 

be labour intensive and may not be appropriate for very large 

invasive species populations. The method is appropriate for 

herbaceous plants that do not easily break from the root and 

for small saplings of woody plants (Holt, 2009). Care should 

be taken while hand-pulling invasive vines to ensure that 

the extensive rooting system is completely removed from the 

soil. Minimize soil or other disturbance to native vegetation. 

[ 13 ]

[ 14 ] 


Figure 3: Removing invasive plants by hand 1 (left) and with a weed wrench 1 (right). 

It is always important to protect yourself by wearing gloves, 

long-sleeved shirts, pants and appropriate footwear. For larger 

shrubs and small trees tools such as weed wrenches may be 

required to pry the roots from the soil (Miller et al. 2010). 

Excavation involves using spades or shovels to remove the whole 

root system. It is only appropriate for very small populations 

because it is very labour intensive and causes a high degree 

of soil disturbance (Miller et al. 2010). Excavation can be an 

effective method of control for plants that break off easily from 

their roots such as dog-strangling vine (Vincetoxicum spp.; see 

section 5.1.10). 

The purpose of flower-head removal is to prevent seed 

production. However, since it is very difficult to ensure that all 

flowers are removed throughout the season, this method should 

only be used to reduce seed production. Flower-head removal 

may be appropriate in sensitive areas where soil disturbance or 

chemical applications are restricted (Frey et al. 2005). 

Cutting can be done using weed whackers, scythes and lawn 

mowers. The objective is to cut the stems as close to the ground 

as possible. Cutting may not kill a plant but it can reduce seed 

production. Cutting right after flowering is most efficient 

because the plant already spent a significant amount of energy 

in the production of flowers. This method generally needs to be 

applied several times throughout the season to prevent any late 

seedpod development. Cutting is appropriate for plants that 

rely on seeds for dispersal (Holt, 2009). 

Mulching involves covering the area requiring control with 

materials such as straw, sawdust or mulched wood and bark. 

This effectively prevents light from reaching germinating 

seedlings. Solarization consists of covering the area with black 

plastic. This generates high temperatures that destroy the soil’s 

seed bank (Holt, 2009)(Fig. 4). 

Directed flame can be an effective treatment for some trees 

and shrubs. It involves applying a directed flame to the base of


Figure 4: Mulching 1 (left) and solarization 1 (right) management techniques. 

the stem using a propane torch. The flame should be applied 

until the base is completely burnt. Several treatments may be 

necessary to kill the invasive plant. This method should only be 

used when the forest floor is wet or damp to decrease the risk 

of fire (Ward et al. 2009; Ward et al. 2010). 

2.3.2 

chemical control 

The use of pesticides in Ontario is highly regulated. The 

Ontario Pesticides Act provides a legal framework for managing 

pesticides that is enforced by the MOE. Pesticides must be 

registered and classified in Ontario. Using an unregistered 

or homemade pesticide is illegal. It is also illegal to use a 

pesticide in any way other than that described by the pesticide 

product label (MOE, 2011). It should be noted that at the time of 

writing not all invasive species in this guidebook have labelled 

pesticides. Please consult with a licenced exterminator before 

attempting any chemical control methods. Refer to the PMRA, 

MOE and/or the OMNR for more information on pesticide use 

in Ontario. 

There are currently 11 classes of pesticides in Ontario (Table 

3). The regulations governing the Pesticides Act were recently 

rewritten to restrict the use of pesticides for cosmetic purposes. 

The purpose of the ban is to reduce public exposure to pesticides 

by preventing their use for aesthetic purposes on residential 

properties and in cemeteries, parks and school yards. Class 5 

or 6 products containing class 11 ingredients are considered 

lower risk pesticides and biopesticides. These are the only 

pesticides that have not been banned for cosmetic uses. Some 

can be used for invasive species control in hardwood stands 

as long as they are being used according to the product label 

(MOE, 2011). 

[ 15 ]

[ 16 ] 


Table 3: Ontario pesticide classification (MOE © Queen’s Printer for Ontario, 2011). 

Class Description Details 

1 Products intended for 

manufacturing purposes. 

2 Restricted or commercial 

products. 


products. 


products. 

5 Domestic products intended 

for household use. 

6 Domestic products intended 

for household use. 

7 Controlled sale products 

(domestic or restricted). 

8 Domestic products that are 

banned for sale and use. 

9 Pesticides are ingredients in 

products for use only under 

exceptions to the ban. 


products for the poisonous 

plant exception. 


products for cosmetic uses 

under the ban. 

Manufacturing concentrates used in the manufacture of a 

pesticide product. 

Very hazardous commercial or restricted pesticides that continue 

to be used under an exception to the ban (i.e. agriculture, forestry, 

golf courses). 

Moderately hazardous commercial or restricted pesticides that 

continue to be used under an exception to the ban (i.e. agriculture, 

forestry, golf courses). 

Less or least hazardous commercial or restricted pesticides that 

continue to be used under an exception to the ban (i.e. agriculture, 

forestry, golf courses). 

Less hazardous domestic pesticides that can be used by 

homeowners and include biopesticides and certain lower risk 

pesticides allowed for cosmetic purposes. 

Least hazardous domestic pesticides that can be used by 

homeowners and include biopesticides and certain lower risk 

pesticides allowed for cosmetic purposes. 

Includes controlled sales domestic pesticides with cosmetic and 

non-cosmetic uses. Products are only allowed to be used for noncosmetic 

purposes. 

Banned domestic products. 

Pesticide ingredients that are banned for cosmetic use. Products 

containing these ingredients may still be used for exceptions to 

the ban. 

Ingredients contained in pesticide products allowed for use under 

the public health or safety exception. These are only ingredients 

that may be used to control plants that are poisonous to the touch 

under the public health or safety exception. 

Ingredients contained in pesticide products that are biopesticides 

or contain lower risk pesticides.

Table 4: Exceptions to the Ontario Pesticide Ban (MOE © Queen’s Printer for Ontario, 2011). 

Exception Applicable circumstances Reasons for exception 

Forestry Treed areas larger than 1 ha. To protect trees from pests and to control 

competing vegetation. 

Arboriculture Upon written recommendation by a 

certified arborist or registered forester. 

Natural 

resources 


In areas where an invasive species is widespread or causing 

a high degree of harm, a class 9 pesticide may be required. 

There are several exceptions to the ban that allow other classes 

of pesticides (e.g., class 9). Relevant exceptions pertaining 

to invasive species in hardwood stands include forestry, 

arboriculture, natural resources and agriculture (Table 4). 

Upon MNR approval. Only relevant when 

no other exception applies. 

To protect tree health. 

To control an invasive species that can pose 

a threat to human health, the environment 

or the economy. 

Agriculture Uses related to agriculture by a farmer. To protect agricultural operations. 

Pesticides are often used to manage insect pests and pathogens 

affecting trees or to control competing vegetation. Class 9 

pesticides can be applied if a hardwood stand is being used for 

forestry activities and if the stand is greater than 1 ha. Under 

the forestry exception, class 9 pesticides can only be applied by 

a person who holds a forestry exterminator licence. Landowners 

may apply for this licence or they may choose to hire a licenced 

exterminator (MOE, 2011). 

If landowners are concerned about a particular tree in their 

woodlot, and if the woodlot is smaller than 1 ha or not being 

used for forestry, they may seek out the help of a tree care 

professional (i.e., a certified arborist or registered forester). If 

the tree care professional determines that the tree is at risk 

unless a pesticide is used he/she may write a letter stating their 

recommendation. The letter will allow the landowner to apply 

a class 5, 6, or 7 product containing a class 9 ingredient to the 

tree. A landscape licenced exterminator can be hired to either 

apply those pesticides or use a class 2, 3, or 4 product (MOE, 

2011). 

In natural areas where the forestry exception does not apply 

(e.g., hardwood stands greater than 1 ha that are not being used 

for forestry activities), a natural resources exception may apply. 

Products containing class 9 ingredients may be used for the 

[ 17 ]

[ 18 ] 


purpose of protecting a native species or its habitat, protecting 

a rare ecosystem or parts thereof, or to control an invasive 

species. Woodlot owners must complete a written application at 

their local MNR office to receive approval before commencing 

chemical control (OMNR, 2010a). 

Woodlot owners who also maintain an agricultural operation 

may use their agriculture class licence. However, these woodlots 

must be associated in some way with the agricultural operation 

in order for the licence to apply. Otherwise a forestry licence 

will be needed to use class 9 pesticides for management in 

the woodlot. Producing crops for use primarily by the owner’s 

household is not considered an agricultural operation (MOE, 

2011). 

herBicide APPlicAtion 

techniques For licenced 

Forestry exterMinAtors 

FoliAr sPrAy 

The foliar spray technique is appropriate for use on herbaceous 

and woody invasive plants. Herbicides are applied directly to the 

foliage using a backpack sprayer or a spray bottle. Herbicides 

should completely cover any new growth on woody plants and 

all the foliage on herbaceous plants. Exterminators should 

completely cover the leaf surface with the herbicide while being 

careful to prevent any runoff. In areas where invasive plants 

intermingle with desirable vegetation a handheld wick or roller 

can be used to directly apply the herbicide to the foliage. This 

eliminates the chance of accidentally spraying the desirable 

vegetation (Miller et al. 2010). The foliar spray technique 

can only be used when leaves are present, thereby limiting 

management to a seasonal timeframe. On the other hand, foliar 

sprays may be used on a variety of both woody and herbaceous 

plants. Therefore, areas with multiple invasive species can be 

treated simultaneously (Swearingen & Pannill, 2009). 

BAsAl BArk 

Basal bark application is appropriate for small diameter trees, 

shrubs and woody vines. Herbicides are applied to the lower 

portion of the stem using a spray bottle or backpack sprayer. 

Depending on the size of the invasive plant, a wide band is 

sprayed around the entire circumference of the stem. Herbicides 

should be applied abundantly so that the entire surface is

coated. Any exposed roots should also receive treatment. Other 

techniques such as streamline and thin line application involve 

applying herbicides in a thin band around the circumference 

of the stem. Streamline application uses diluted herbicides 

whereas thin line uses non-diluted herbicides appropriate for 

small diameter stems (Miller et al. 2010). Basal bark application 

works best in the late winter or early spring when most native 

plants are still dormant. The stem needs to be completely dry 

and free of ice and snow for the herbicide to work effectively 

(Swearingen & Pannill, 2009). 

steM injection 

Trees, shrubs and large woody vines can be treated using the 

stem injection technique. Cuts are made in the stem diagonally 

towards the ground creating a small crevice through which the 

herbicide is poured. These cuts should be made around the entire 

stem and placed approximately 2.5cm apart (Miller et al. 2010). 

A spray bottle can be used to fill the crevice with just enough 

herbicide to prevent dripping. Herbicides must be applied to 

the cuts immediately, before the plant begins sealing off the 

wounded area. Herbicide applicators are available in various 

forms. Blades, drills and injection systems can be purchased. 

These applicators cause wounds in the bark through which 

herbicides are inserted simultaneously (Swearingen & Pannill, 

2009). Caution should be taken when using stem injection 

techniques as some species may exude certain herbicides and 

translocation may affect neighbouring trees, thereby increasing 

the risk of non-target hardwood mortality (Lewis & McCarthy, 

2008). 

cut-stuMP 


Some managers may decide to first cut and remove invasive 

trees, shrubs or woody vines from the forest stand. Although 

this method may be labour intensive and time consuming it 

may be practical in areas where large infestations make access 

to the woodlot difficult. Investing time and energy in areas 

with dense invasions will ultimately make future management 

easier (Swearingen & Pannill, 2009). After the above-ground 

portion of the stem is removed, herbicides can be applied 

directly to the stump using spray bottles, brushes, handheld 

wicks or rollers. Most herbicides should be applied immediately 

after cutting, and with any sawdust removed, to allow for 

optimal absorption. Large invasions may need to be divided 

into smaller units so that cutting and herbicide application 

can be accomplished on the same day. For stumps larger than 

7.5cm in diameter herbicides should be applied to wet the outer 

edge of the cut surface while the cut surface of smaller stumps 

should be completely covered (Miller et al. 2010). 

[ 19 ]

[ 20 ] 


3.0 

A triBute to hArdwood Forests 

oF ontArio 

3.1 

timber harvest 

3.1.1 

ontario’s Forests 

Ontario has a total area of 107.6 million ha, of which 66% is 

forested. The majority of these forests, approximately 81%, 

is Crown land owned by the province (Fig. 5). A total of 27.1 

million ha fall within the Area of Undertaking (AOU) where 

forest management activities actively take place. The majority of 

privately owned forests occur in southern Ontario and account 

for approximately 10% of the forested land in Ontario. A total of 

9% are protected in parks and recreational areas (OMNR, 2012a). 

There are four forest regions in Ontario, including the deciduous 

forest, the Great Lakes-St. Lawrence forest, the boreal forest and 

the Hudson Bay lowlands (Fig. 6). The occurrence of these forest 

regions is due to a combination of differences in bedrock, soil 

type and climate (Armson, 2001). 

The deciduous forest region on the southern tip of Ontario is 

the most biologically diverse of the province’s forest regions. It 

is unique to Ontario, not being found anywhere else in Canada 

(Armson, 2001). Most was converted to agricultural land during 

European settlement. Today this area is highly populated and 

the majority of forested land occurs in scattered woodlots, 

parks and conservation areas (OMNR, 2012a).

Crown - Outside AOU 

Crown (AOU) 

Protected Area 

Private 

Federal/First Nations 

A TRIBUTE TO HARDWOOD FORESTS OF ONTARIO 3.0 

Figure 5: Land ownership classes in Ontario 64. Figure 6: Ontario’s forest regions 64. 

The Great Lakes-St. Lawrence forest comprises a mixture of 

hardwood and softwood species. It is the second largest forest 

region in Ontario and occurs in both populated and rural areas 

(OMNR, 2012a). Both the deciduous forest and Great Lakes- 

St. Lawrence forest regions occur within the Great Lakes-St. 

Lawrence lowlands, which is characterized by sedimentary 

rock, level topography and relatively fertile soils (Wake et al. 

1997). 

The boreal forest and Hudson Bay lowlands lie within the 

northern latitudes of Ontario. The boreal forest region comprises 

mainly softwood species. It is the largest forest region in the 

province, containing 58% of Ontario’s forested land (OMNR, 

2012a). The boreal coincides with much of the Canadian Shield 

where thin soils and pre-Cambrian rock prevail (Wake et al. 

1997). The Hudson Bay lowlands occur along the northern edge 

of Ontario. Spruce, tamarack, willow and birch dominate this 

forest region. Other prominent species are trembling aspen 

and balsam poplar (Armson, 2001). Areas of forested land are 

intermingled with large areas of open muskeg and the landscape 

is dotted with small ponds and lakes. This forest region occurs 

over sedimentary bedrock (OMNR, 2012a). 

Hudson Bay 

Lowlands 

Boreal Forest 

Great Lakes-St. 

Lawrence Forest 

Deciduous 

Forest 

[ 21 ]

[ 22 ] 


3.1.2 

history, economy and culture of 

timber harvest 

Historically, wood exports have played an important role in 

the Canadian economy. The wood trade in Ontario was a direct 

result of European demand. When the regular exports of wood 

from the Baltic region to Britain were compromised due to the 

Napoleonic wars, Britain was left in desperate need of wood 

for naval construction (Lower, 1933). As a result, demand 

for softwood timber more than doubled in the early 1800’s 

(Armson, 2001). 

The demand for wood products created a form of livelihood 

that allowed for the immigration of people to Upper Canada, 

now known as Ontario. British demand called for squared 

timber, which required logs to be cut on four sides. The method 

was wasteful, leaving behind 25 to 30% of the tree (Easterbrook 

& Aitken, 1988). However, square timber was in demand and 

accounted for over 50% of Canadian exports (Bothwell, 1986). As 

wood was slowly replaced by iron and steel in construction, the 

European demand for timber began to decrease. At this time, 

a new market was opening up with the United States where the 

demand for sawn logs was rising (Easterbrook & Aitken, 1988). 

Historically, softwoods were the major commodity of the timber 

trade. White pine was the most solicited species followed by 

red pine and white spruce. Hardwood from maple and oak was 

also desirable, albeit to a smaller extent than softwood (Lower, 

1933). Large, straight softwood trees were primarily targeted in 

forest stands since square timber greater than 20cm in width 

was in demand. As softwood species such as white pine became 

depleted, the industry shifted to hardwood exports; a result of 

a change in supply instead of demand (Armson, 2001). 

Today, timber products continue to be an important part of 

Ontario’s economy. Currently there are over 200 000 forestryrelated 

jobs in the province, many in communities dependant 

on the industry. In 2008, forestry products made $14 billion in 

revenue for Ontario; the majority coming from pulp and paper 

products (MNDM, 2011).


3.2 

Maple syrup Production 

3.2.1 

history 

the oriGins oF MAPle syruP 

The exact origin of maple syrup is unknown. European settlers 

like to claim the discovery; however, there is evidence for sugarmaking 

being long-established as part of the native traditions. 

The Algonquin word Sinzibuckwud (maple sugar) or the words 

Sisibaskwatattik (maple tree) and Sisbaskwat (sugar) of the Cree 

tribe have no similarities to the European settler’s language. It 

stands to reason that there would be some similarities if the 

settlers had crafted those terms (Nearing & Nearing, 1950). The 

Cree did adopt a word for the white sugar that settlers brought 

with them. The word Sukaw has been said to be an attempt at 

pronouncing either the English word ‘sugar’ or the French word 

‘sucre’ (Henshaw, 1890). There are also many early writings by 

settlers that mention the native people’s previous knowledge 

of tapping maple trees and making maple syrup. The Nearings 

(1950) point out an example found in a letter written by Robert 

J. Thornton which was published in London’s Philosophical 

Magazine, 1798: 

“I have enclosed some sugar of the first boiling got from the juice 

of the wounded maple…Twas sent from Canada, where the natives 

prepare it from said juice; eight pints yielding commonly a pound 

of sugar. The Indians have practiced it time out of mind…” 

(Thornton cited in Vogel, 1987). 

There are several native legends that relate to the discovery of 

maple syrup. One talks of a time when maple syrup ran thick 

and pure from maple trees. After discovering this, villagers got 

lazy and started spending their days drinking syrup instead 

of hunting and fishing. A mischief-maker, known by names 

such as Glooskap and Ne-naw-bo-zhoo, sees that the villagers 

are taking the syrup for granted. He fills the trees with water 

from the river so as to dilute the sap. From that day forward, 

villagers have been required to boil down the sap to enjoy the 

pure form of maple syrup (Chamberlain, 1891). 

[ 23 ]

[ 24 ] 


Another legend tells of a chief’s wife. The chief decides to go 

hunting for the day and grabs his tomahawk from the maple 

tree he had left it in the night before. Throughout the day sap 

leaks from the wound and collects in a basket that happens to 

be at the base of the tree. In a rush to get some water boiling 

for dinner, the chief’s wife grabs the bucket full of sap and 

proceeds to cook their evening meal. The food flavoured with 

maple was greatly enjoyed and the discovery shared with all 

(Lawrence & Martin, 1993). 

Ininatigs’s gift of sugar is another story. It tells of a family’s 

hardship during a long, cold winter. Food supplies had been 

depleted and the family was on the verge of starvation. From 

out of the forest comes a voice. It is the voice of Ininatig, the 

man tree. He tells the family to cut him and carefully collect 

the liquid flowing from his wounds. By boiling this liquid the 

family could make syrup and sugar that would sustain them 

and save their lives (Wittstock, 1993). 

The final legend involves a hungry hunter who is watching 

the antics of a red squirrel in a grove of maple trees. The 

squirrel seemed to be biting holes in the branches and eating 

the emerging sap. The hunter may have collected some sap 

and heated it, possibly multiple times, to sustain him through 

a period of hunger (Lawrence & Martin, 1993). A study by 

Heinrich (1992) shows that red squirrels (Tamiasciurus 

hudsonicus) ‘tap’ sugar maple trees. They bite on the branches 

creating small wounds. Observations show that red squirrels 

do not eat the sap right away but wait for some of the water to 

evaporate and concentrate the sugar. Perhaps the maple tree’s 

secret was discovered by someone who decided to try an icicle. 

The sap running from broken branches generally freezes to 

produce these icicles. During the freezing process, some water 

evaporates leaving a vaguely sweet taste (Hauser, 1998). 

the evolution oF MAPle syruP 

Production 

First Nation people used to tap maple trees by creating a 

V-shaped wound in the tree. A piece of wood or bark was placed 

at the bottom of the wound to allow the sap to flow away from 

the tree into a wooden container placed at the base (Lawrence 

& Martin, 1993). Birch bark containers or wooden pots were 

used for heating the sap. As these types of containers could 

not withstand fire, hot stones were dropped into the containers 

as a means of heating the sap. Stones were replaced as needed 

until the sap boiled down to the desired consistency. Another 

method used was freezing. Sap would be left overnight in 

shallow containers to freeze and the resulting layer of ice that


formed on top would be taken away, leaving the concentrated 

syrup underneath (Nearing & Nearing, 1950). 

The iron kettle came with the arrival of European settlers 

(Fig. 7). These kettles greatly increased the quality and quantity 

of sap that could be produced (Whitney & Upmeyer, 2004). The 

use of multiple kettles was eventually adopted for boiling sap. 

As the sap achieved a desired consistency it could be transferred 

to the next kettle and so forth until it reached a finished state. 

This method helped to prevent burning and off-setting the 

flavour of the syrup (Nearing & Nearing, 1950). 

Trees continued to be tapped with the traditional V-shaped slash 

for up to 80 years after the arrival of European settlers. The 

creation of these large wounds would significantly harm the tree 

and usually decrease its sap producing capacity considerably 

(Whitney & Upmeyer, 2004). Augers eventually replaced the 

use of axes to bore less harmful holes in the trees. Although 

techniques such as cutting branches and roots to collect sap 

were attempted, the trunks have remained the traditional 

location for tapping. Buckets were eventually attached to trees 

as a means to prevent high winds from blowing the dripping 

sap out of the buckets’ range (Nearing & Nearing, 1950) (Fig. 8). 

Figure 7: Kettles for boiling sap 3. 

Figure 8: A bucket for collecting sap 2. 

[ 25 ]

[ 26 ] 


Collection and transportation of sap was labour intensive. Sugar 

shacks were virtually non-existent and most boiling occurred 

out in the sugar bush. Most maple syrup producers would 

plan to spend the entire season out in their sugar bush. Small 

shelters would occasionally be constructed to protect the fire 

and boiling sap from the elements (Fig. 9). A central fire would 

be constructed and sap transported by hand or with horse and 

sleigh to this central location (Nearing & Nearing, 1950). 

Figure 9: A sugar shack at night 2. 

Modern MAPle syruP Production 

Maple syrup producers in Ontario must meet the requirements 

of Regulation 386 of the Farm Products Grades and Sales Act 

if they are going to sell their products locally. To sell outside 

of Ontario, producers must comply with the Maple Products 

Regulations found in the Canada Agricultural Act (Chapeskie 

& Hendersen, 2007). The CFIA is the governing body for maple 

syrup production. This agency is responsible for ensuring that 

maple syrup producers make and sell both a safe and high 

quality product. 

The quality of maple syrup in Ontario is governed by a threetier 

grading system which is based on colour and taste. The 

first grade, Canada No. 1, describes a category of syrup ranging


in colour from extra light to medium (Heiligmann et al. 2006). 

This grade of maple syrup has a mild flavour and is often used 

as a condiment. Syrup that gets a Canada No. 2 grade has an 

amber colour class. It is often used in cooking due to its strong 

flavour (Chapeskie & Hendersen, 2007). Canada No. 3 is a dark 

syrup and the maple flavour may be off-set by a caramel or 

sappy taste. Only pure maple syrup, with a sugar content of at 

least 66%, can be graded in Canada (Heiligmann et al. 2006). 

A product’s colour class is assigned depending on the amount 

of light that can pass through it. The grade is given according 

to flavour. Commonly lighter syrups have a very mild taste 

whereas darker syrups have a strong candied flavour. 

Soil characteristics, timing and handling all contribute to 

differences in colour and taste among syrups. Most producers 

agree that sap collected earlier in the season will produce a 

higher quality product lighter in colour and milder in flavour 

compared to that made from sap collected later in the season 

(Hortvet, 1904). 

Today there is a wide variety of technology to help alleviate the 

work and cost associated with maple syrup production while 

ensuring a high quality product. Plastic tubing has greatly 

decreased labour costs required for transporting sap from 

trees to the holding tank (Lawrence & Martin, 1993) (Fig. 10). Sap 

can move through the tubing by gravity or it can be pumped 

through. Vacuum pumps were adopted as a means of increasing 

sap production during times of imperfect weather conditions 

when sap flow is slow (Coons, 1992). Some producers continue 

to use buckets for sap collection but usually only those with 

small operations. Buckets must be collected daily to prevent 

microbial growth and labourers are generally friends or family 

members (Lawrence & Martin, 1993). 

Figure 10: Sap lines 1 (left) and buckets 3 (right). 

[ 27 ]

[ 28 ] 


The adoption of sheet metal was a turning point in the history 

of maple syrup production. Metal replaced wooden spouts 

and pails. The use of large flat-bottomed pans replaced the 

old iron kettles. After much experimenting with these pans 

a primitive evaporator began to emerge (Lawrence & Martin, 

1993). The modern-day evaporator is essentially a large, twochambered 

pan (Fig. 11). The first chamber has a ridged bottom 

that maximizes the surface area for heating sap quickly and 

efficiently. Sap is directed through the chamber in a serpentine 

pattern. The final chamber has a flat bottom to prevent the 

sap from burning. When the syrup has reached the desired 

consistency it is passed through a filter and left to cool. Reverse 

osmosis machines were adapted to help concentrate the sap 

before boiling. These machines helped to reduce boiling time, 

thereby saving on energy costs associated with the long hours 

required for syrup production. They separate water from the 

soluble sugars found in maple sap through a membrane and by 

working on a pressure gradient (Heiligmann et al. 2006). 

Figure 11: Evolution of flat-bottomed pans 2 to modern day evaporators 3. 

3.2.2 

economic and cultural 

significance 

econoMic vAlue 

Maple syrup is only produced in Canada and the northern part 

of the United States. This is due to the relationship between sap 

flow and weather conditions. Canada is the leading producer of 

maple syrup with the majority of production coming from four 

provinces: Quebec, Ontario, New Brunswick and Nova Scotia. 

Ontario has approximately 2600 maple syrup producers (Leuty, 

2009).


Ideal weather conditions in 2009 resulted in a record year for 

the maple syrup industry, with a total revenue of $353.8 million 

in Canada. Of this amount Ontario contributed $25.6 million. In 

2008, Canada totaled $211.9 million with Ontario contributing 

$15.4 million (Statistics Canada, 2009). 

The ability to produce maple sugar was of great value to the 

early settlers. It was too costly to import sugar and many 

families had to do without. The timing of the sap run was 

ideal for the farmer because it would not overlap with the crop 

growing season. Time could be spent collecting and boiling 

large amounts of sap for syrup and sugar production. After 

enough product was made and set aside for family use the rest 

could be sold for a profit (Whitney & Upmeyer, 2004). 

Maple syrup production is not the necessity it once was for 

settlers. However, it is still an important product in Ontario. 

The economic value of maple syrup production benefits rural 

communities. It allows rural families to create an income and 

it contributes to tourism. Indeed, maple syrup is embedded 

in the image of Canada and represents a major asset for 

branding the country’s tourism, food and agricultural sectors 

internationally. A trip to the sugar shack in the spring is a 

long-standing tradition in many families. The rustic feel of a 

traditionally run sugaring operation is a major attraction to 

the tourist (Hinrichs, 1995). From the viewpoint of a producer, 

making maple syrup allows one to make use of the land while 

developing a rural livelihood (Hinrichs, 1998). 

heritAGe vAlue 

The maple leaf is meaningful for Canadians as individuals 

(Fig. 12). The maple leaf appears not only on the National 

Flag of Canada but also on the Canadian penny and on both 

the Ontario and Quebec coat of arms. In 1996 the maple tree 

officially became Canada’s national arboreal emblem. It is no 

wonder that Canadians hold a sentimental place for the maple 

tree and its products when its leaf holds such an integral part 

of the country’s identity (DCH, 2008). 

Maple syrup production is a traditional part of Canada’s heritage 

(Fig. 13). It is one of the earliest agricultural activities to exist 

in Ontario (Coons, 1992). It is associated with festivities, renews 

family bonds and brings communities together. The sugar 

season occurs in the spring and represents a time of renewal, a 

time to leave behind the feelings of isolation that a cold winter 

can create and a time of gathering to renew friendships. Perhaps 

it is the fact that syrup operations were, and still commonly 

are, family run businesses that lends romance to the industry. 

[ 29 ]

[ 30 ] 


Figure 12: A national emblem 3. 

Running a sugar bush represents a lifestyle that many long to 

experience – even if only for a day at the sugar shack (Whitney 

& Upmeyer, 2004). 

Many maple syrup producers in Ontario sell their product 

directly to the consumer. In today’s society, maple syrup is no 

longer the sole source of sugar in a household. It is considered 

a luxury item. Buying syrup directly from the vendor holds an 

appeal. It allows one to briefly connect with the past and celebrate 

the Canadian heritage. Many operators tend to shy away from 

an overabundance of new technology. They want to keep their 

operations looking traditional because it attracts customers 

who want to experience their rural heritage (Hinrichs, 1995). 

Figure 13: Creating family traditions 2.

4.0 

woodlot MAnAGeMent 

Managing a hardwood stand has many benefits for the landowner 

or woodlot manager. Whether the goal is to manage for timber 

harvest, maple syrup production or nature conservation, it is 

important to become familiar with the age of the forest and 

species composition. This can be done by completing a forest 

inventory. Forest inventories can be done by the landowner or 

by a hired consultant. The following two sections outline some 

basic management concepts depending on whether the goal of 

the woodlot is for timber harvest or maple syrup production. 

Managing a woodlot helps to keep the forest healthy. In turn, a 

healthy forest is generally less susceptible to invasion by exotic 

plants, insects and disease (Hilts & Mitchell, 1999). 

4.1 

Management for timber harvest 

Written by Fraser Smith, M.Sc., Forestry, Ontario Federation of Anglers and Hunters 

Stand structure describes the distribution of tree ages and 

size classes within a forest stand (Chapeskie et al. 2006). Stand 

structure helps to determine what has occurred in a stand 

previously, and what silvicultural actions should take place 

to best manage the forest for optimal health and long-term 

productivity. Broadly, there are two major categories of stand 

structure: even-aged and uneven-aged. Even-aged forests are 

composed of trees that are all similar in age, usually all within 

10 years. Even-aged forests often experience stand-replacing 

4.0 

[ 31 ]

[ 32 ] 


disturbance patterns such as fires, after which most trees 

regenerate. This forest type is more typical of softwood stands, 

but can also be seen in the sun-loving pioneer species such as 

aspen (Populus spp.) and birch (Betula spp.). Managers of evenaged 

forests strive to open the canopy to allow sufficient light to 

reach these shade intolerant species and promote regeneration 

(Lane, 2007). 

Uneven-aged forests have a broad range of size classes and ages. 

Generally, these forests are prone to small-scale disturbance 

patterns that affect individual or small groups of trees. They 

are composed of shade-tolerant or mid-tolerant species that 

regenerate in heavy or partial shade. Ideally, these forests have 

a large number of juvenile trees, a medium amount of middlesized 

trees, and a small number of very large, old trees. This 

age distribution is characterized by a decreasing number of 

stems with increasing age and size. It fosters the development 

of vigorous young saplings that compete for light and grow 

straight and tall (Lane, 2007). 

Selection harvesting is a management system where individual 

trees or small groups of trees are cut in intervals of 8 to 20 

years. Selection management is designed to mimic the small, 

frequent disturbances such as wind, ice storms, lightning, 

and disease that are characteristic of the hardwood forests 

of southern Ontario (Burke et al. 2011). Each cut may harvest 

up to one third of the stand while maintaining a largely intact 

canopy that promotes regeneration of tolerant to mid-tolerant 

species. The aim of selection harvest is to maintain an unevenaged 

stand with a diverse range of species and stand structure. 

This range of species and structure is most appropriate for 

mid-tolerant hardwood stands and promotes stand health while 

maintaining productivity (Lane, 2007). 

The shelterwood system describes a management approach 

where a mature forest is harvested in two or more passes over 

a period of 5 to 60 years (Burke et al. 2011). Each of these cuts 

or “thinning” passes aims to carefully open up the canopy 

of mature trees and provide the increased space and light in 

the understory for natural regeneration of mid-tolerant trees 

to thrive (OMNR, 2004). The shelterwood system mimics 

disturbance patterns that affect only part of a stand such as 

ground fires, insects, and disease. The shelterwood system is 

most appropriate for species adapted to those disturbances 

such as white pine (Pinus strobus), white spruce (Picea glauca) 

and red oak (Quercus rubra). First, a preparatory cut thins the 

stand but leaves large vigorous trees with the ability to produce 

a large productive crown. Next, roughly 20 years after the 

preparatory cut, the regeneration cut removes approximately 

half of the originally retained trees. This cut allows light to 

reach the forest floor and for seedling regeneration to increase. 

Once regeneration has reached an appropriate level (usually 3 

to 15 years later), one or two removal cuts are performed to

WOODLOT MANAGEMENT 4.0 

harvest some of the remaining mature trees. This gives the 

now established saplings full sunlight and encourages vigorous 

growth (Burke et al. 2011). 

The shelterwood system is most often applied in white pine 

forests, however it is sometimes applied in tolerant hardwood 

forests to promote turnover from a heavily degraded site. While 

the shelterwood system is considered even-aged, there are 

generally two to three distinct age classes growing together. 

These distinct age classes, or cohorts, represent those trees 

established during one of the thinning cuts and are intended 

to follow those natural disturbance patterns found in these 

stands. The shelterwood system should only be carried out 

using an appropriate prescription and employing a certified 

tree marker (OMNR, 2004). 

The clearcut system describes an intensive management 

strategy where the majority of a stand of mature trees is 

harvested in one pass and forest regeneration is initiated in a 

short time-frame. This system is suited to those areas where 

the most common disturbance patterns are stand-replacing 

such as fire, flood, and large windstorms. The dominant tree 

species in Ontario’s boreal forest, such as black spruce (Picea 

mariana), Jack pine (Pinus banksiana), and aspen (Populus spp.), 

regenerate best under the open conditions created by these 

disturbances. Since these species require full sunlight without 

competing shade, the clearcut system is designed to emulate 

these natural disturbance patterns and facilitate rapid stand 

renewal. If these stands were to be harvested using a selection 

or shelterwood system, the structure and species composition 

of this forest would be fundamentally altered from the original 

forest. This in turn would lead to cascading effects for the 

ecology of the area, including wildlife that depend on these 

stands and their natural replacement. In Ontario, clearcut 

systems never remove all trees on a site. Mature trees are left 

either scattered individually across the site (called clearcut 

with seed trees) or in small patches (aggregated structural 

retention). Regeneration of these sites can be either natural or 

through planting (Burke et al. 2011). 

Whatever reason a landowner has for harvesting their woodlot, 

keeping site damage to a minimum should always be a primary 

goal. Site damage from harvest activities can include residual tree 

injury, soil compaction and rutting, and introductions of exotic 

invasive plant species that can have lasting implications for the 

future of the stand. Keep in mind that disturbance intensity 

appears to be positively correlated with a forest’s susceptibility 

to the establishment and spread of invasive species (Bell & 

Newmaster, 2002). To avoid harvest-related damage, all woodlot 

operations should adhere to careful logging practices. For a 

complete guide to minimizing harvest damage, the Ontario 

Woodlot Association has produced “A Landowner’s Guide to 

Careful Logging” (Byford, 2009) which provides landowners 

[ 33 ]

[ 34 ] 


with sound advice as they make decisions to protect the health 

and integrity of their woodlots. Landowners planning harvest 

operations should also be aware of the potential for invasive 

species introductions. Invasive species can be accidentally 

transported as seeds, plant fragments or in soil on recreational 

or harvest equipment. Every attempt should be made to clean 

equipment before transport, minimize soil disturbance, and 

follow careful logging practices. 

4.2 

sugar Bush Management 

Management helps to increase the health and productivity of a 

sugar bush. Trees in a forest are in constant competition with 

one another for light, space, water and nutrients. This creates 

stress on maple trees and does not allow for optimum growth. 

For example, trees that are in competition for light will put 

more energy into growing tall so as to reach the canopy. As a 

consequence, these trees remain small in diameter and have 

less potential for reaching an adequate size to be useful for 

sap collection (Richardson, 2003). Trees with large crowns are 

desired because they produce a larger quantity of sweet (i.e., 

more concentrated) sap as compared to maples with small 

crowns. Crown development in sugar maples is a function of 

space and light availability (Chapeskie et al. 2006). Management 

allows for improved growth of sugar maple crowns through 

thinning procedures. Certain trees are harvested from the 

sugar bush to ensure that adequate resources are available for 

crop trees, which will grow both faster and larger. Trees with 

larger diameters will also produce more sap because they can 

withstand a greater number of taps (Coons, 1992). Management 

enables a maple syrup producer to tap a higher number of trees 

in a smaller amount of time (Richardson, 2003). 

Management depends on the size and age class of the forest. 

A smaller forest with 300 taps is managed differently than a 

forest with 10 000 taps. Thinning a small sugar bush must 

be carried out very carefully. If too many trees are removed 

the sugar bush will lose taps and decrease in productivity. 

Likewise, an even-aged forest would be managed differently 

than a forest containing a variety of age classes. In a sugar 

bush where all the trees are roughly the same age, regeneration 

is a primary concern since there will be no new trees to replace 

the old ones for quite some time. A sugar bush containing trees 

of a variety of ages is generally thought to be ideal because 

there will always be younger trees available to replace the old 

(Richardson, 2003).

WOODLOT MANAGEMENT 4.0 

The goal of sugar bush management should be to sustain a 

healthy forest while keeping the future in mind. Focusing only 

on today’s sap production could shorten the life of a sugar 

bush considerably. Thus, one main component of management 

is regeneration. Maple trees are long-living but they will not 

produce an adequate amount of sap consistently and across the 

entire life span of a tree. New trees will eventually be required 

to replace the old. One problem with regeneration has to do 

with the amount of time it takes for a sugar maple to reach 

a size where it can be tapped. Seedlings in the understory 

require a certain amount of space and sunlight to grow. Thus, 

management practices should consider the undergrowth 

(Coons, 1992). 

There is an ongoing debate as to whether diversity or 

monoculture is the better management strategy for a healthy 

sugar bush. In the past, managers have been advised to harvest 

all species, other than maple, in the sugar bush. This strategy 

would free up more space for maple trees to grow and create 

a good environment for sugar maple saplings to take hold. 

Managers are now being advised that a diversity of species 

makes a healthier sugar bush because monocultures are more 

susceptible to insect outbreaks and disease. Managers must 

decide: by allowing the growth of other species there will be less 

crop trees in the forest and the production may be smaller than 

in a monoculture. On the other hand, going with a monoculture 

increases the risk of insect and disease outbreaks (Lawrence & 

Martin, 1993). 

There are many benefits associated with maintaining a diversity 

of trees in a sugar bush. Biodiversity enhances a range of 

ecological functions within a forest. It creates wildlife habitat 

that can support predators of sugar maple pests. Some sites 

within the sugar bush may not be suitable for maple growth. 

By allowing other species to grow in these areas there is the 

added benefit of a periodic timber harvest. Wood can be sold 

for a profit or used to fuel the evaporators. A diverse forest has 

also been known to be more resistant and resilient to insect 

outbreaks (Chapeskie et al, 2006). 

Management generally includes a forest inventory and a 

management plan. A forest inventory includes a list of species, 

age and diameter of individual trees, and the overall number of 

trees (OMNR, 2006). The relative abundances of tree species are 

recorded, allowing the manager to determine what proportion 

of the forest comprises maple and the general biodiversity of 

the area. Tree diameter and overall health are recorded for 

individual species greater than 10cm in diameter. Density 

and basal area are obtained to determine whether thinning is 

required to reduce the amount of competition around maple 

trees (Chapeskie et al. 2006). The inventory allows the producer 

to make informed decisions when it comes to making the 

management plan. Monitoring should be an ongoing part of 

[ 35 ]

[ 36 ] 


management. It allows the producer to deal with problems, 

such as damage due to invasive species as soon as they arise 

(OMNR, 2006). 

To aid producers with management decisions such as thinning, 

stocking recommendations are available. Table 5 lists the 

recommended number of trees per hectare based on average 

trunk diameter. It also gives the producer an idea as to the 

average number of taps a sugar bush of a particular diameter 

class can support (Richardson, 2003). As stated earlier, the 

goal of management is to produce a more sustainable forest, 

with healthy and productive maple trees. It is important to 

understand that improper tapping can have negative effects on 

trees and sap production. For example, trees that are stressed 

from other causes such as drought may not be able to withstand 

the additional stress caused by tapping. The number of taps per 

tree should depend on the diameter and overall health of each 

individual tree (Richardson, 2004). 

Table 5: Sugar bush stocking and tapping recommendations (adapted from 

Richardson, 2003; 2004). 

Tapping rule 

Normal 

(healthy trees) 

Conservative 

(unhealthy trees) 

Average 

diameter (cm) 

Number 

of taps 

Recommended 

trees/hectare 

Number of 

taps/hectare 

< 10 0 > 680 0 

10 – 25 0 210 – 680 0 

26 – 37 1 150 – 209 150 – 209 

38 – 50 2 100 – 149 200 – 298 

51 – 63 3 66 – 99 198 – 297 

≥ 64 4 < 66 < 364 

30-46 1 NA NA 

≥ 47 2 NA NA 

Management should be adjusted to provide the greatest amount 

of revenue both in the short and long term. It must be flexible 

to account for unpredictable events such as severe weather or 

invasive species outbreaks, which can reduce the sugar bush’s 

productivity (Chapeskie et al. 2006). The following chapters on 

invasive species will cover best management practices, early 

detection techniques and prevention strategies to alleviate the 

negative effects associated with invasive species.

5.0 Invasive Species 

Accounts 

5.1 Exotic Plants 

5.2 Exotic Insects and 

Pathogens

[ 38 ] 


5.1 

exotic Plants 

5.1.1 Norway Maple (Acer platanoides) HIGH 

5.1.2 Tree-of-heaven (Ailanthus altissima) HIGH 

5.1.3 Garlic Mustard (Alliaria petiolata) HIGH 

5.1.4 Barberry (Berberis thunbergii & B. vulgaris) HIGH 

5.1.5 Japanese Knotweed (Fallopia japonica) HIGH 

5.1.6 English Ivy (Hedera helix) HIGH 

5.1.7 Himalayan Balsam (Impatiens glandulifera) HIGH 

5.1.8 Common Buckthorn (Rhamnus cathartica) HIGH 

5.1.9 Periwinkle (Vinca minor) HIGH 

5.1.10 Dog-strangling Vine (Vincetoxicum rossicum & V. nigrum) HIGH 

5.1.11 Goutweed (Aegopodium podagraria) MODERATE 

5.1.12 Oriental Bittersweet (Celastrus orbiculatus) MODERATE 

5.1.13 Exotic Bush Honeysuckle (Lonicera spp.) MODERATE 

5.1.14 Kudzu (Pueraria montana var. lobata) MODERATE

5.0 INVASIVE SPECIES ACCOUNTS 

5.1.1 

norway Maple 


Other common names: 

schwedler maple, crimson king maple 

Priority Rating: hiGh 

Figure 14: Norway maple 1. 

39 

[ ]

[ 40 ] 


identiFicAtion 

leAves: Norway maple has dark green leaves with an opposite 

arrangement (Fig. 15). Each leaf has 5 pointed lobes (Kaufman 

& Kaufman, 2007). The tops of the leaves are a darker green 

as compared to the lighter, shiny undersides. A milky sap is 

emitted from leaf stalks when they are cut (Farrar, 1995). 

Figure 15: Norway maple leaves 1 (left) and opposite leaf arrangement 1 (right). 

Flowers: Yellow-green flowers grow in clusters at the end of 

branches (Fig. 16). Each flower has 5 petals and 5 sepals. The 

flowers appear at the same time as the leaves emerge in early 

spring (Kershaw, 2001). 

Fruit/seeds: The fruit, also called a key, consists of a winged 

seedcase with an angle of 180° between the wings (Fig. 17). Two 

seeds are developed per fruit. The wings are approximately 35 

to 50mm long and the seedcases have a flattened appearance 

(Farrar, 1995) 

Figure 16: Norway maple flowers 4 . Figure 17: The fruit of Norway maple 5 .

5.1.1 NORWAY MAPLE (Acer platanoides) 

trunk: Norway maples are 

large trees that range in height 

from 12 to 21m (Kaufman 

& Kaufman, 2007; Kershaw, 

2001). The bark of mature 

trees has deep grooves with 

interlacing ridges (Fig. 18). The 

bark is dark gray and lenticels 

are often prominent on young 

branches (Farrar, 1995). 

siMilAr sPecies 

sugar maple (Acer saccharum) 

Figure 18: The bark of Norway 

maple 6 . 

Sugar maple is a large shade tree that is very similar in 

appearance to Norway maple. There are several differences 

between the leaves and the keys that can help with identification. 

The leaves of Norway maple are of a much darker green than 

the yellow-green leaves of sugar maple (Fig. 19). The leaves of 

Norway maple only turn yellow in the fall whereas sugar maple 

leaves range in colour from yellow, orange and red. The angle 

between the wings of sugar maple keys is 120° whereas the 

angle between the wings of Norway maple keys is 180° (Farrar, 

1995; Kershaw, 2001) (Fig. 20). 

Figure 19: Comparing a sugar maple leaf 1 (left) to that of Norway maple 1 (right). 

Figure 20: Comparing sugar maple keys 5 (left) to those of Norway maple 7 (right). 

[ 41 ]

[ 42 ] 


Black maple (Acer nigrum) 

Black maple is a large shade tree. Generally the leaves of black 

maple have 3 lobes but they may have 5 lobes at times (Fig. 21). 

The smooth edged leaves with 5 lobes may appear very similar 

to those of Norway maples. However, the leaf undersides and 

stalks of black maple are hairy. Figure 22 shows how the keys of 

black maple have wings that are parallel to one another whereas 

those of Norway maple have a distinct 180° angle (Farrar, 1995; 

Kershaw, 2001). 

Figure 21: Comparing black maple leaves 1 (left) to those of Norway maple 1 (right). 

Figure 22: Comparing black maple keys 5 (left) to those of Norway maple 8 (right). 

red maple (Acer rubrum) 

Red maple is a medium-sized shade tree. Red maple leaves have 

3 to 5 lobes with many small, irregular teeth along the edges. 

The notches on either side of the central lobe have a distinctive 

V-shape whereas those of Norway maple have a gently curving 

U-shape (Fig. 23). The keys of red maple have an angle of 60° 

between the wings, whereas the wings of Norway maple keys 

are separated by an angle of 180° (Farrar, 1995; Kershaw, 2001) 

(Fig. 24).


Figure 23: Comparing a red maple leaf 1 (left) to that of Norway maple 1 (right). 

Figure 24: Comparing the keys of red maple 7 (left) to those of Norway maple 6 (right). 

silver maple (Acer saccharinum) 

Silver maple is a medium-sized shade tree. Silver maple leaves 

have 5 to 7 lobes that are distinctly separated by deep notches. 

The leaf edges have irregular teeth whereas the leaf edges of 

Norway maple are smooth (Fig. 25). Silver maple keys have an 

angle of 90° between the wings, which is much narrower than 

that found on Norway maple wings (Farrar, 1995; Kershaw, 

2001) (Fig. 26). 

Figure 25: Comparing a silver maple leaf 1 (left) to that of Norway maple 1 (right). 

[ 43 ]

[ 44 ] 


Figure 26: Comparing the keys of silver maple 5 (left) to those of Norway maple 4 (right). 

striped maple (Acer pensylvanicum) 

Striped maple is a shrub or a small understory tree that only 

reaches heights of 10m. The leaves have 3 lobes and regular 

toothed edges whereas Norway maple has leaves with 5 lobes 

and smooth edges (Fig. 27). Figure 28 shows the differences 

between the keys of striped maple and Norway maple. Notice 

that the keys of striped maple only have a 90° angle between 

wings as opposed to the 180° angle found in Norway maple keys 

(Farrar, 1995; Kershaw, 2001). 

Figure 27: Comparing a striped maple leaf 9 (left) to that of Norway maple 1 (right). 

Figure 28: Comparing the keys of striped maple 10 (left) to those of Norway maple 4 (right).


Mountain maple (Acer spicatum) 

Mountain maple is a shrub or a small understory tree that only 

grows about 5m tall. Mountain maple leaves may have a similar 

5-lobed appearance to those of Norway maple. However, the 

edges are distinct. Mountain maple has irregular toothed edges 

whereas Norway maple has smooth leaf edges (Fig. 29). There 

is less than a 90° angle between the wings of mountain maple 

keys, while Norway maple keys have an angle of 180° between 

wings (Farrar, 1995; Kershaw, 2001) (Fig. 30). 

Figure 29: Comparing a mountain maple leaf 1 (left) to that of Norway maple 1 (right). 

Figure 30: Comparing the keys of mountain maple 1 (left) to those of Norway maple 7 (right). 

[ 45 ]

[ 46 ] 


A key to plants that may be confused with 

norway maple (Acer platanoides) 

1...Leaves.with.toothed.edges 

. 2..Leaves.with.deep.lobes.(5-7.lobes)...........................................................................Silver.maple.(Acer saccharinum) 

. 2..Leaves.with.shallow.lobes.(3-5.lobes) 

. . 3..Single-toothed.leaves.with.teeth.curved.on.one.side...................................Mountain.maple.(Acer spicatum) 

. . 3..Irregular,.double-toothed.leaves.with.teeth.straight.on.both.sides.................. 

. . . 4..Finely.pointed.and.uniform.teeth....................................................................Striped.maple.(Acer pensylvanicum) 

. . . 4..Sharply.pointed,.irregular.teeth.......................................................................Red.maple.(Acer rubrum) 

1..Leaves.with.smooth.edges 

. . . . 5..Leaf.undersides.and.stalks.hairy..................................................................Black.maple.(Acer nigrum) 

. . . . 5..Leaf.undersides.and.stalks.hairless 

. . . . . 6..Yellow-green.leaves.with.3-5.bluntly.pointed.lobes.......................Sugar.maple.(Acer saccharum) 

. . . . . 6..Dark.green.leaves.with.5-7.sharply.pointed.lobes............................Norway.maple.(Acer platanoides). 

tAxonoMic hierArchy 

Kingdom. . . . . . . Plantae 

. Subkingdom. . . . . Tracheobionta 

. . Division. . . . . . Magnoliophyta 

. . . Class. . . . . . Magnoliopsida 

. . . . Subclass.. . . Rosidea 

. . . . . Order. . . . Sapindales 

. . . . . . Family.. . Sapindaceae 

. . . . . . . Genus.. Acer 

. . . . . . . . Species. Acer platanoides 

BioloGy 

oriGin & distriBution 

Norway maple is native to Europe where it is 

widespread and can be found growing naturally 

in mixed forests. Due to its popularity as a 

street tree in Europe it was only a matter of 

time before Norway maple became available for 

purchase as an ornamental in North America. 

It is thought to have first been introduced in 

1756 in Philadelphia, Pennsylvania. Today, 

Norway maple can be found in six Canadian 

provinces, including Ontario, Quebec, Prince 

Edward Island, New Brunswick, Newfoundland 

and British Columbia (Nowak & Rowntree, 

1990). 

hABitAt 

Norway maple grows well in both open and 

closed habitats. It is a popular street tree and 

is commonly found in urban areas. Due to its


shade tolerance, Norway maple has spread into natural areas 

and can be found anywhere from urban woodlots to relatively 

undisturbed forests (Fig. 31). It grows better in mesic soils than 

in dry or very wet areas (Bertin et al. 2005; Meiners, 2005). 

Figure 31: Fall colours of Norway maple in a forest 6 . 

reProduction 

Norway maple’s sexual reproduction drives its spread. The 

flowers are insect-pollinated and seeds are produced in large 

quantities. Seeds germinate readily to form extensive seedling 

banks. These seedlings wait in the understory until an opening 

is created in the canopy (Fig. 32). As sunlight penetrates to the 

seedlings below, they can quickly grow in height to fill the 

available space (Webb et al. 2001). 

Figure 32: Norway maple seedlings in a forest understory 6 . 

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[ 48 ] 


liFe cycle 

Norway maple flowers early in the season, usually in April and 

May as the leaves expand (Nowak & Rowntree, 1990). Fruits are 

produced in the summer and mature in the autumn. Leaves 

senesce and turn yellow a couple of weeks later than native 

maples. Some winged fruits may stay on the tree throughout 

the winter (Kershaw, 2001). 

success MechAnisMs 

Norway maple has a longer growing season as compared to 

native maples. Its leaves expand earlier and senesce later, 

thereby extending the growing season by as much as 3 weeks 

(Kloeppel & Abrams, 1995). This extra growing time may allow 

Norway maple to produce a larger and hardier seed crop. A 

superior seed crop allows for greater capacity to regenerate. 

The seeds germinate into shade tolerant seedlings that thrive 

in the understory, waiting for suitable conditions to grow into 

the canopy. Whenever light becomes available through an 

opening in the canopy the seedlings of Norway maple have the 

ability to rapidly grow in height. This rapid growth rate gives 

Norway maple a competitive advantage when it comes to canopy 

recruitment (Martin, 1999). 

Not only does Norway maple produce a large quantity of seeds 

but each individual seed is hardy. Norway maple seeds are 

double and sometimes triple the weight of sugar maple seeds. 

A larger seed mass may benefit Norway maple by increasing its 

chances of survival and germination in a shaded understory. In 

fact, germination rates were observed to be higher for Norway 

maple as compared to sugar maple (Martin & Marks, 2006). 

Norway maple may have an advantage in North America due to 

a lack of herbivores. One study documented less leaf and fungal 

damage to Norway maples in North America as compared to 

those found in Europe (Adams et al. 2009). Resources normally 

needed to repair damage done by herbivores can be put 

towards growth and reproduction. Even though Norway maple 

experiences some herbivory in its introduced range, studies 

have shown it to be more resilient to severe insect damage than 

native maples. Foliar insect damage was found to be greater for 

sugar maples as compared to co-occurring Norway maples in 

forests in New Jersey and Pennsylvania (Cincotta et al. 2009).


ecosysteM iMPActs 

Norway maple invasions can suppress native hardwood 

regeneration (Fig. 33). In a 5ha woodlot in New York, concerns 

were raised that sugar maple was being replaced by Norway 

maple. Martin (1999) found that sugar maple (Acer saccharum) 

saplings were absent under Norway maple canopy trees. Norway 

maple saplings were found in greater abundance relative to 

other hardwood species and more than half of the saplings 

growing under sugar maple canopies were Norway maples. 

Figure 33: Invasion of Norway maple saplings 6 . 

Similar effects on hardwood regeneration were observed in an 

18ha forest preserve in New Jersey. Norway maple regeneration, 

in relation to total number of saplings, was found to be double 

that of American beech (Fagus grandifolia) and over five times 

that of sugar maple (Webb & Kaunzinger, 1993). Moreover, when 

Norway maple was present in a mixed oak forest in central New 

Jersey, the survival and growth of red maple (Acer rubrum), 

white elm (Ulmus americana) and red oak (Quercus rubra) were 

significantly reduced (Galbraith-Kent & Handel, 2008). 

Norway maple invasions may inhibit the growth and occurrence 

of native wildflowers and shrubs. Norway maple casts a deeper 

shade as compared to native forest trees, which may eliminate 

or reduce shade intolerant species. Also, Norway maple’s prolific 

reproduction allows for high seedling densities, which impart 

a high competitive pressure on other understory species. In a 

New Jersey forest preserve, fewer species were found growing 

under Norway maple canopies than under native sugar maple 

(A. saccharum) and American beech (F. grandifolia). In fact, 

Norway maple recruits occupied the majority of space under all 

three canopies (Wyckoff & Webb, 1996). 

[ 49 ]

[ 50 ] 


vectors & PAthwAys 

Norway maple can easily escape 

cultivation as its windborne seeds 

can disperse long distances. At 

times, seeds can disperse over 100 

metres away from the parent tree 

(Bertin et al. 2005). Since Norway 

maple is commonly planted as a 

street tree (Fig. 34) many areas may 

be exposed to its seeds, especially 

urban woodlots (Anderson, 1999). 

Small mammals feed on Norway 

maple seeds and may contribute to 

its dispersal (Meiners, 2005). 

MAnAGeMent PrActices 


control oPtions 

Figure 34: Norway maple in an urban centre 6 . 

• Use native maples in horticulture plantings; 

• Remove any Norway maples from the property, especially 

large individuals that are capable of producing a large 

quantity of seeds; 

• Minimize soil disturbance whenever possible. 


• Learn how to properly identify Norway maple and be able to 

easily distinguish it from native maples; 

• Monitor the woodlot frequently, paying particular attention 

to disturbed areas or places with recent openings within the 

canopy; 

• Survey your local area for any large Norway maple trees that 

could be a potential seed source. 

Hand-pulling: Small seedlings can be hand-pulled and weed 

wrenches can be used for saplings less than 3cm in diameter 

(Strobl & Bland, 2000). Soil disturbance should be minimized 

while pulling because it may promote the recruitment of new


Norway maple seedlings. Removal of small individuals in the 

understory should coincide with the removal of all mature 

seed-producing trees in the vicinity. Management efforts in the 

understory will eventually exhaust the seed bank and allow 

native species, such as sugar maple, to regenerate (Webb et al. 

2001). 

Cutting and Girdling: Cutting and girdling can be an effective 

non-chemical means of control for trees and saplings larger 

than 3cm in diameter. Cutting mature trees will eliminate the 

seed source and ease the burden of future management efforts. 

However, cutting down large trees will create an opening in the 

forest canopy, which may promote the growth of Norway maple 

saplings. To minimize this problem, ensure that management of 

seedlings and saplings in the understory coincide with canopy 

tree management. Divide large areas into manageable sections 

for control. Another option is to girdle large trees. It may take 

a few growing seasons for the tree to die. However, there will 

be no sudden openings in the canopy to encourage understory 

growth. Cut any re-sprouts from stumps as required (Webb et 

al. 2001; Clauson, 2011). 

Herbicide application: Chemical control is effective in controlling 

Norway maple trees and to prevent re-sprouting. The cut- 

stump, basal bark and hack-and-squirt methods are all effective 

(Webster et al. 2006). Foliar application for understory saplings 

is generally not advised. Norway maple often grows in close 

proximity to sugar maple and other desirable hardwood species. 

Small seedlings and saplings can be hand-pulled. Alternatively, 

a cut-stump method can be used in conjunction with herbicide 

application to the cut surface of the stem after the saplings 

have been removed (Swearingen et al. 2010). Refer to section 

2.3.2 for explanation of chemical application methods. 

recoMMendAtions For inteGrAted 

control oF lArGe invAsions 

option #1: with chemical control 

Target large seed producing trees first by applying herbicides 

using the basal bark, hack-and-squirt or cut-stump methods. 

The basal bark and hack-and-squirt methods may be easier than 

the cut-stump method in terms of labour as they do not require 

large trees to be felled. However, large dead standing trees may 

pose a hazard in some areas and may need to be felled anyway. 

Keep in mind that the cut-stump method will create large 

canopy openings that may promote invasion by other exotics 

in the vicinity. Monitoring such areas and controlling where 

needed, will give native species such as sugar maple time to fill 

in the canopy gaps (Swearingen et al. 2010). 

[ 51 ]

[ 52 ] 


After all seed producing trees are removed, focus can be put 

on the understory. Young seedlings under 3cm in diameter can 

be pulled. Anything larger can be cut followed by a herbicide 

application to the cut stem to prevent any re-sprouting. Norway 

maple often grows in the same areas as native sugar maple. 

Thus, applying herbicides using a foliar spray may cause 

undesirable damage (Strobl & Bland, 2000). 

option #2: without chemical control 

Large seed producing trees should be cut or girdled to prevent 

additions to the seed bank. Cut and girdled trees will likely resprout 

and will thus require frequent clipping. After all large 

trees have been controlled, management efforts can focus on 

smaller seedlings and saplings. As Norway maple takes several 

years to mature before seed production, understory control 

efforts can be scheduled every couple of years. Norway maple 

responds well to soil disturbance which is a natural result of 

management. Thus, allowing native saplings time to grow and 

become established may promote healthy competition with 

native plants (Galbraith-Kent & Handel, 2008). 

Table 6: Management recommendations for Norway maple (Acer platanoides). 

Extent.of.infestation Small.invasions.and.satellite. 

populations 

Recommended. 

method.of.control 

Hand-pulling.seedlings.and.young. 

saplings..Cutting.trees.that.are.too. 

large.to.be.hand-pulled. 

Large.invasions.and.dense.populations 

Integrated.control: 

(Combination.of.physical.and.chemical. 

control). 

Timing Spring,.summer.and.fall. Cutting.and.hand-pulling.can.be.done. 

any.time.in.the.spring,.summer.and.fall.. 

Herbicide.application.should.occur.in.the. 

spring.or.fall.while.other.native.plants.are. 

dormant. 

Frequency.of.control Cut.trees.and.saplings.will.likely.resprout.and.require.frequent.clipping. 

Length.of.control 

2-3.years. 5+.years. 

Required.restoration Hand-pulling.creates.soil.disturbance. 

that.benefits.Norway.maple. 

recruitment..Consider.planting.native. 

vegetation.or.transplanting.with.sugar. 

maple.seedlings. 

Cut.trees.and.saplings.will.likely.re-sprout. 

and.require.frequent.clipping..Plan.to. 

control.Norway.maple.in.the.understory. 

every.couple.of.years. 

Consider.planting.native.vegetation.in. 

disturbed.areas..Large.dead.standing.trees. 

may.become.a.hazard.and.may.need.to.be. 

removed.


5.1.2 

tree-of-heaven 

(Ailanthus altissima) 


varnish tree, stinktree, chinese sumac, Ailanthus, copal tree, 

stinking sumac 


Figure 35: Tree-of-heaven 11 . 

53 

[ ]

[ 54 ] 



leAves: Tree-of-heaven has alternately arranged compound 

leaves. The compound leaves are composed of 11 to 41 leaflets 

(Fig. 36). Each leaflet has 1 to 4 glandular lobes near the base 

(Hu, 1979; Farrar, 1995). 

Figure 36: The compound leaves 12 (left) and a single leaflet 1 (right) of tree-of-heaven. 

Flowers: Tree-of-heaven is dioecious (i.e., each individual 

tree has either male or female flowers). These yellow-green 

flowers grow in large clusters called panicles (Fig. 37). Male 

trees generally produce more flowers, forming panicles that 

are 3 to 4 times larger than those found on female trees (Hu, 

1979). 

Figure 37: Tree-of-heaven flower panicles 4 (left) and individual flowers 4 (right).

5.1.2 TREE-OF-HEAVEN (Ailanthus altissima) 

Fruit/seeds: Seeds are produced in winged fruits called 

samaras, each holding a single seed (Fig. 38). The samaras form 

dense clusters, with a mature tree producing over 300 000 

seeds. Only female trees bear fruit (Bory & Clair-Maczulajtys, 

1980). 

Figure 38: Dense samara clusters 4 (left) and individual fruits 11 (right). 

trunk: Tree-of-heaven is a fast growing, medium-sized tree. 

It can increase in height by 2 to 3m per season and range from 

15 to 25m tall (Kershaw, 2001). The trunks of young trees are 

smooth and greenish-grey in colour (Fig. 39). As trees mature 

the bark turns a deeper grey and forms shallow, pale crevices 

(Farrar, 1995). Young branches are hairy and green, turning 

reddish-brown with age. Leaf scars are large and heart-shaped 

(Kowarik & Säumel, 2007). 

Figure 39: Shallow crevices in mature bark 6 (left) and a heart-shaped leaf scar 1 (right). 

[ 55 ]

[ 56 ] 



Black locust (Robinia pseudoacacia) 

Black locust has compound leaves with 7 to 19 leaflets that 

could be confused with those of tree-of-heaven. The leaflets 

of black locust have smooth edges whereas those of tree-ofheaven 

are lobed. Black locust leaves lack the glandular lobe at 

the base of the leaflet characteristic of tree-of-heaven (Fig. 40). 

Pairs of spines located along the stem may also help to identify 

black locust (Fig. 41). After being introduced from the eastern 

United States, black locust has become naturalized over much 

of southern Ontario (Farrar, 1995; Kershaw, 2001). 

Figure 40: Comparing the compound leaves of black locust 5 (left) to 

those of tree-of-heaven 14 (right). 

Ash (Fraxinus spp.) 

Figure 41: The characteristic spines 

of black locust 15 . 

Several native species of ash in Ontario may be mistaken for 

tree-of-heaven. Ash trees have smaller compound leaves, with 

5 to 11 leaflets, compared to the 11 to 41 leaflets of tree-ofheaven 

(Fig. 42). Another distinction is the arrangement of the 

compound leaves along the stem. Ash leaves have an opposite 

arrangement whereas tree-of-heaven displays an alternate 

arrangement (Farrar, 1995; Kershaw, 2001). 

Figure 42: Comparing the compound leaves of black ash 8 (left) to those of tree-of-heaven 16 (right).


Poison-sumac (Toxicodendron vernix) 

Poison-sumac is native to North America. It is a shrub or 

small tree with alternately arranged compound leaves. Each 

compound leaf has 7 to 13 leaflets with smooth edges whereas 

tree-of-heaven has 11 to 41 leaflets with glandular lobes close 

to the base (Fig. 43). Do not touch or burn poison-sumac as its 

oils and smoke can cause skin, eye and lung irritation (Farrar, 

1995; Kershaw, 2001). 

Figure 43: Comparing the compound leaves of poison-sumac 14 (left) to those of tree-of-heaven 5 (right). 

staghorn and smooth sumac (Rhus typhina & R. 

glabra) 

Staghorn and smooth sumac are native to Ontario. They do not 

belong to the same genus as poison-sumac and since they are 

safe to touch they are commonly planted as ornamentals. These 

species of sumac have a similar number of leaflets (11 to 31) per 

compound leaf as compared to tree-of-heaven (11 to 41). However, 

the leaflets of staghorn and smooth sumac are sharply toothed 

unlike those of tree-of-heaven which are smooth with lobed 

bases (Fig. 44). Smooth sumac has hairless branches whereas 

those of staghorn sumac are covered in hair to the point where 

they appear fuzzy. Sumacs emit a characteristic milky sap when 

the leaf stalks or twigs are cut (Farrar, 1995; Kershaw, 2001). 

Figure 44: Comparing the leaflets of staghorn sumac 5 (left) to those of tree-of-heaven 16 (right). 

[ 57 ]

[ 58 ] 


Mountain-ash (Sorbus spp.) 

There are three species of mountain-ash in Ontario. American 

mountain-ash (Sorbus americana) and showy mountain-ash (S. 

decora) are native to Ontario while European mountain-ash (S. 

aucuparia) is exotic. They grow as shrubs or small trees and 

have pinnate, compound leaves consisting of 9-17 leaflets (Fig. 

45). They can be distinguished from tree-of-heaven by the finely 

toothed edges of their leaflets (Farrar, 1995; Kershaw, 2001). 

Figure 45: Comparing the compound leaves of mountain-ash 1 (left) to 

those of tree-of-heaven 6 (right). 

hickory (Carya spp.) 

There are several species of hickory native to Ontario. Hickories 

have compound leaves with 5-11 leaflets (Fig. 46). The terminal 

leaflet is generally larger than the lateral leaflets. Hickory’s 

leaflets have toothed edges while tree-of-heaven’s leaflets have 

smooth, lobed edges (Farrar, 1995; Kershaw, 2001). 

Figure 46: Comparing the compound leaves of hickory 5 (left) to those of tree-of-heaven 16 (right).


Butternut (Juglans cinerea) 

Butternut is a medium-sized tree reaching 12 to 25m in height. 

Its compound leaves have 11 to 17 leaflets with toothed edges 

and are hairy underneath. The top three terminal leaflets are 

equal in size while the rest get progressively smaller from top 

to bottom along the central stalk (Fig. 47). Tree-of-heaven has 

leaflets that are generally equal in size with smooth lobed edges 

(Farrar, 1995; Kershaw, 2001). Butternut is native to Ontario and 

is currently considered a species at risk (OMNR, 2012b). 

Figure 47: Comparing the compound leaves of butternut 8 (left) to those of tree-of-heaven 5 (right). 

Black walnut (Juglans nigra) 

Black walnut is a medium-sized tree, growing up to 30m tall. 

Black walnut is native to Ontario and is prized for its high 

value wood. The compound leaves have 14 to 22 leaflets with 

finely toothed edges and slightly hairy undersides (Fig. 48). 

The terminal leaflet is either absent or much smaller than the 

lateral leaflets. Tree-of-heaven’s top and lateral leaflets are 

generally similar in size (Farrar, 1995; Kershaw, 2001). 

Figure 48: Comparing the compound leaves of black walnut 1 (left) to those of tree-of-heaven 11 (right). 

[ 59 ]

[ 60 ] 



tree-of-heaven (Ailanthus altissima) 

1..Branches.with.spines.............................................................................................................................Black.locust. 

. . . . . . . . . . (Robinia pseudoacacia) 

1..Branches.without.spines 

. 2..Opposite.compound.leaves............................................................................................................Ash.(Fraxinus spp.) 

. 2..Alternate.compound.leaves 

. . 3..Leaflets.with.smooth.edges........................................................................................................Poison-sumac 

. . . . . . . . . . (Toxicodendron vernix) 

. . 3..Leaflets.with.toothed.or.lobed.edges 

. . . 4..Leaflets.with.sharp.regular.teeth 

. . . . 5..Leaf.stalks.and.twigs.release.milky.sap.when.cut 

. . . . . 6..Branches.very.hairy.........................................................................................................Staghorn.sumac. 

. . . . . . . . . . (Rhus typhina) 

. . . . . 6..Branches.hairless…..........................................................................................................Smooth.sumac.(Rhus glabra) 

. . . . 5..Leaf.stalks.and.twigs.do.not.release.a.milky.sap.when.cut 

. . . . . . 7..Leaflets.on.a.compound.leaf.are.all.similar.in.size 

. . . . . . . 8..Leaflets.with.hairy.undersides............................................................................European.mountain-ash 

. . . . . . . . . . (Sorbus aucuparia) 

. . . . . . . 8..Leaflets.with.smooth.undersides 

. . . . . . . . 9..Leaflets.3-5.times.as.long.as.they.are.wide.with.a.tapering.point......American.mountain-ash 

. . . . . . . . . . (Sorbus americana) 

. . . . . . . . 9..Leaflets.2-3.times.as.long.as.they.are.wide.with.an.abrupt.point.......Showy.mountain-ash. 

. . . . . . . . . . (Sorbus decora). 

. . . . . . 7..Leaflets.on.a.compound.leaf.are.dissimilar.in.size 

. . . . . . . . . 10..Terminal.leaflet.larger.than.adjacent.leaflets......................................Hickory.(Carya spp.) 

. . . . . . . . . 10..Terminal.leaflet.equal.in.size.or.smaller.than.adjacent.leaflets 

. . . . . . . . . . 11..Three.terminal.leaflets.equal.in.size...................................................Butternut.(Juglans cinerea) 

. . . . . . . . . . 11..Terminal.leaflet.absent.or.smaller.than.the.rest.............................Black.walnut.(Juglans nigra) 

. . . 4..Leaflets.with.lobes.near.the.base.........................................................................................Tree-of-heaven. 

. . . . . . . . . . (Ailanthus altissima)


Kingdom. Plantae. 

..Subkingdom. Tracheobionta 

....Division. Magnoliophyta 

.....Class. Magnoliopsida 

......Subclass. Rosidae 

.......Order. Sapindales 

........Family. Simaroubaceae 

.........Genus. Ailanthus 

..........Species. Ailanthus altissima 


BioloGy 


Tree-of-heaven is native to China. 

Seeds were supposedly shipped 

to Europe and planted in Paris in 

the early 1740’s. In 1784 it was first 

introduced to North America in a 

garden in Philadelphia. Tree-of-heaven 

was highly valued as an ornamental 

plant due to its rapid growth and lush 

foliage. As a result, it was commonly 

planted in urban areas where it readily 

escaped cultivation (Fig. 49). Tree-ofheaven 

is found in southern Ontario 

but has the potential to expand north 

(Hu, 1979). 

Figure 49: Tree-of-heaven growing alongside a building 15 . 

[ 61 ]

[ 62 ] 


hABitAt 

Tree-of-heaven is shade intolerant and has historically been 

documented to invade open, disturbed areas. It is considered a 

nuisance in urban areas where it grows close to buildings and 

through the cracks in sidewalks (Hu, 1979). Due to its shade 

intolerance, tree-of-heaven was not traditionally considered 

a threat to forests. However, recent studies have documented 

tree-of-heaven invasions in old- and second-growth forests 

(Kowarik, 1995; Knapp & Canham, 2000) (Fig. 50). 

Figure 50: Tree-of-heaven occupying space in the canopy 11 . 

reProduction 

Tree-of-heaven is dioecious (i.e., each tree is either male or 

female). Seeds are produced in winged fruits on female trees 

(Hu, 1979). A mature tree can produce more than 300 000 seeds


(Bory & Clair-Maczulajtys, 1980). Although tree-of-heaven 

reproduces mainly through seeds, it can also reproduce 

asexually by way of clones that readily sprout from the root 

and stem (Kowarik & Säumel, 2007). 

liFe cycle 

Tree-of-heaven takes 3 to 5 years to mature. Only after this 

period will trees produce flowers and, consequently, produce 

seeds. Flowering occurs from mid-April through June and 

samaras are produced from August to October. These ripen 

from September until the end of October and germinate the 

following year (Kowarik & Säumel, 2007). In one study, a small 

percentage of seeds remained viable for a second growing 

season, implying that tree-of-heaven may establish a shortterm 

seed bank (Kota et al. 2007). Further investigation into the 

longevity of such a seed bank is required. 


Tree-of-heaven has the advantage of being able to reproduce 

sexually and asexually. A single tree can produce more than 

300 000 seeds that are easily dispersed by wind (Bory & Clair- 

Maczulajtys, 1980) (Fig. 51). Tree-of-heaven’s fast growth rate 

and preference for light allows it to quickly occupy gaps 

in the forest canopy created by management activities or 

natural causes (Knapp & Canham, 2000). Once canopy trees 

are established their root systems can branch out into the 

understory. Nutrients transferred from the parent to saplings 

emerging from its roots alleviate the stress imposed by the lowlight 

environment (Kowarik, 1995). 

Figure 51: Prolific seed production by tree-of-heaven 4 . 

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[ 64 ] 


Tree-of-heaven synthesizes secondary compounds that can be 

found in both the living tissue and the surrounding soil (Lawrence 

et al. 1991). These compounds can inhibit the establishment 

and growth of other plant species (Mergen, 1959; Heisey, 1990; 

Lawrence et al. 1991) and act as a defense against herbivores 

and pathogens (Heisey, 1990), thereby strongly enhancing treeof-heaven’s 

competitive ability against native species. 

ecosysteM iMPActs 

Tree-of-heaven may inhibit native hardwood regeneration 

through allelopathic effects and direct competition. Such effects 

could result in a shift in the species composition of the forest 

and thus change the dynamics of the forest community (Gómez- 

Aparicio & Canham, 2008b). Once established, tree-of-heaven 

can form dense patches (Fig. 52) and, due to its fast growth rate, 

directly compete with native vegetation for nutrients and canopy 

space (Knapp & Canham, 2000). 

Figure 52: A dense population of tree-of-heaven 15 . 


Seeds are contained in winged capsules (samaras) that enable 

long distance, windborne dispersal. Seeds can easily be spread 

by wind currents into the interior of the forest from parent trees 

located at the forest edge (Landenberger et al. 2007). Water is 

another agent of dispersal for seeds and plant fragments that 

can produce clones (Kowarik & Säumel, 2008). Rodents may also 

help to disperse seeds by collecting and caching them for the 

winter (Kowarik & Säumel, 2007).




• Do not dump yard wastes in natural areas. Seeds or stem fragments 

may be incorporated in the wastes and could easily germinate or 

re-sprout in refuse piles; 

• Refrain from planting tree-of-heaven as an ornamental on your 

lawn. Seeds can disperse long distances and could easily enter a 

nearby woodlot or natural area. 


• Learn how to properly identify tree-of-heaven at all life stages. 

Young saplings can easily be confused with other native understory 

species (Fig. 53). Tree-of-heaven is most easily identified by the 

characteristic lobes at the base of the leaflets (see above section on 

similar species); 

• Monitor the woodlot frequently and pay particular attention to 

disturbed areas and forest edges. 

Figure 53: Tree-of-heaven saplings 1 (left) are easily confused with sumacs 19 (right). 


Hand-pulling: Hand-pulling is only practical for small seedlings. 

Larger saplings develop a taproot that is difficult to remove 

by hand (Hoshovsky, 1995). Although the use of tools such as 

a weed wrench may help to remove larger saplings, they can 

break the root system, leaving behind root fragments that can 

re-sprout. Moist soils make pulling easier and increase the 

likelihood that the entire root system will come loose. Clonal 

shoots arising from trailing roots may be difficult to remove 

as they are attached to other individuals. Moreover, removing 

the root system without leaving behind root fragments is nearly 

[ 65 ]

[ 66 ] 


impossible. Care should be taken to remove as much of the 

root as possible to prevent re-sprouting (Swearingen & Pannill, 

2009). 

Cutting and girdling: Cutting and girdling are often ineffective 

means of control for tree-of-heaven because it can promote resprouting 

and make the invasion worse (Meloche & Murphy, 

2006). Girdling will kill large trees, however, the roots will 

send up new clonal shoots. Cutting and girdling will only 

make the problem worse if frequent ongoing management is 

not an option. This is because frequent cutting will eventually 

exhaust the plant’s reserves and kill the tree. To be successful, 

this method will require several years, with frequent repeated 

cuttings per year (Burch & Zedaker, 2003). 

Excavation: Large trees and saplings can be cut and their entire 

root system excavated from the ground. However, digging out 

these stumps and roots is labour intensive and causes a great 

deal of soil disturbance, thus promoting re-invasion. Excavation 

is only practical in areas where there are only a few individuals. 

After management, holes should be filled and replanted with 

native vegetation or covered with mulch to help decrease the 

chance of re-establishment by tree-of-heaven and other exotics 

(Swearingen & Pannill, 2009). 

Herbicide application: Herbicide application is an effective 

way of controlling tree-of-heaven invasions. It can help to kill 

existing trees while effectively preventing re-sprouting that 

is difficult to avoid with other non-chemical control methods 

(Burch & Zedaker, 2003). It is best to use a systemic herbicide 

when trying to control tree-of-heaven because they are 

absorbed by the plant tissues and transported below-ground to 

the entire root system. This is important because clonal shoots 

can continue to sprout even if the above-ground portion of the 

parent tree is dead. Use hack-and-squirt or cut-stump methods 

of herbicide application to large trees. Saplings may be treated 

using the cut-stump or basal bark method. Seedlings and 

saplings may be treated using a foliar spray (Miller et al. 2010). 




Tree-of-heaven is often difficult to eradicate and control. Its 

abundant seed production coupled with vegetative spread 

makes frequent monitoring a necessity. In heavily invaded 

areas it may be more practical to target mature female trees, 

which can produce over 300 000 seeds per year. Once the large 

female trees have been removed, management efforts can focus 

on the male trees and the smaller saplings in the understory. 

Several years of monitoring may be required to catch any re-


sprouts or germinating saplings from a potentially short-term 

seed bank (Miller et al. 2010). 

Small seedlings can easily be pulled before they develop a taproot 

whereas larger saplings may either be pulled by using a 

root wrench, excavation of their root systems or through a 

foliar herbicide application. In the case of mature trees it is 

important to target the female seed-producing trees first to 

reduce the addition of seeds into the area. The hack-and-squirt 

method may be applied to the large trees. Alternatively, the 

cut-stump method can be used keeping in mind the additional 

labour requirements of cutting and removing large trees from 

the area. In all situations, frequent monitoring will be needed to 

control any re-sprouts. A second application of herbicides may 

be needed to kill large trees or “stubborn” saplings (Miller et al. 

2010). 


Controlling large populations of tree-of-heaven without using 

herbicides will require repeated monitoring and management. 

Targeting large seed-bearing female trees will reduce the 

number of seedlings emerging the following year. Cutting these 

large trees will promote re-sprouting from the stump and roots. 

However, if these re-sprouts are frequently cut back the root 

reserves will eventually be exhausted. The initial tree felling 

should occur in the early summer, just after flowering, to 

minimize the proliferation of seeds. In addition, the root system 

is most vulnerable at this time because most of the energy 

has been spent on producing leaves and flowers. Promoting a 

healthy overstory that enhances the amount of shade will also 

discourage re-invasion (Pannill, 2000). 

After all the large female trees have been removed, the invaded 

area should be split up into manageable sizes for control. Areas 

along the outer boundary of the invasion should be targeted 

first, working towards the centre of the most heavily invaded 

area. Large trees should be cut while small saplings should 

be pulled by hand. To completely destroy root reserves and 

prevent re-sprouting repeated control measures will be needed 

throughout the season and for multiple years (Hoshovsky, 1995). 

The above-ground portion of tree-of-heaven should be put 

through a wood chipper or burned. For small invasions with 

a few individual trees a manager might choose to leave the 

slash as woody debris to enhance wildlife habitat. However 

caution should be taken due to tree-of-heaven’s potential for 

allelopathy. In heavily invaded areas or where it could interfere 

with management activities, the wood can be turned into 

woodchips or firewood. Large slash piles may also be burned on 

site. Since even small root fragments can sprout into a new tree, 

the roots of tree-of-heaven should be carefully disposed of. It is 

recommended to bag or incinerate the roots (Hoshovsky, 1995). 

[ 67 ]

[ 68 ] 


Extent.of. 

infestation 

Recommended. 


Table 7: Management recommendations for tree-of-heaven (Ailanthus altissima). 

Small.invasions.and.satellite.populations Large.invasions.and.dense.populations 

Hand-pulling,.excavation.and.cutting. Integrated.control.including.hand-pulling,. 

cutting.and..herbicide.application. 

Timing Hand-pull.all.emerging.saplings.in.the. 

spring.before.they.develop.a.large.taproot. 

Cut.and.excavate.larger.saplings. 

throughout.the.year. 

Cut.large.trees.in.the.early.summer.right. 

after.flowering.for.optimal.results. 

Frequency.of. 

control 

Ongoing.monitoring.and.control.will.be. 

needed..Re-sprouts.from.cut.trees.will. 

need.to.be.controlled.several.times.per. 

year. 

Length.of.control 2-5.years. 5+.years. 

Required. 

restoration 

Plant.native.species.or.apply.mulch.in. 

areas.where.hand-pulling.creates.soil. 

disturbance. 

Hand-pull.all.emerging.saplings.in.the. 

spring.before.they.develop.a.large.taproot. 

Cut.large.trees.in.the.early.summer.right. 

after.flowering. 

Apply.herbicides.in.the.spring.or.fall.when. 

most.native.vegetation.is.dormant. 

Ongoing.monitoring.and.management. 

will.be.needed.to.control.tree-of-heaven. 

invasions..Re-sprouts.from.cut.trees.will. 

need.to.be.controlled.several.times.per. 

year..Herbicides.may.need.to.be.reapplied.the.following.year. 

Plant.native.species.or.apply.mulch.in. 

areas.where.hand-pulling.creates.soil. 

disturbance.


5.1.3 

Garlic Mustard 



hedge Garlic 


Figure 54: Garlic Mustard 1. 

69 

[ ]

[ 70 ] 



Growth ForM: Garlic mustard has a two-year life cycle. 

During the first year, a small cluster of leaves called a basal 

rosette form (Fig. 55). During the second year, stems emerge 

from the basal rosette to produce flowers and seed. Flowering 

stems can reach 1.25m in height. All parts of the plant emit a 

garlic odour when crushed (Cavers et al. 1979). 

Figure 55: First-year basal rosette 1 (left) and second-year growth 1 (right) of garlic mustard. 

leAves: Basal rosette leaves are kidney-shaped with wavy 

edges (Rodgers et al. 2008). Deep veins give the leaves a wrinkled 

appearance. Each rosette has 2 to 12 leaves arranged in a whorl. 

These leaves remain green throughout the winter, even when 

buried in snow (Fig. 56). Second-year leaves grow from a stalk in 

an alternate arrangement (Fig. 57). They are triangular in shape 

with toothed edges (Cavers et al. 1979). 

Figure 56: First-year basal leaves of garlic mustard 1 . Figure 57: Second-year leaves of garlic mustard 1 .

5.1.3 GARLIC MUSTARD (Alliaria petiolata) 

Flowers: Flowers have 4 white petals, 4 green sepals and 6 

stamen (Fig. 58). They grow in clusters, also known as racemes, 

at the top of each stem (Cavers et al. 1979). 

Figure 58: Garlic mustard flowers 1 . 

Fruit/seed: Mature plants produce long, slender seedpods 

full of tiny black seeds (Kaufman & Kaufman, 2007) (Fig. 59). 

Figure 59: Seedpod formation 1 (left) and mature seeds 1 (right) of garlic mustard. 

root: Taproots have an S-shaped pattern (Fig. 60) directly 

below the stem (Cavers et al. 1979). 

Figure 60: Characteristic S-curved root of garlic mustard 1 . 

[ 71 ]

[ 72 ] 



Ground ivy (Glechoma hederacea) 

The first-year basal leaves of garlic mustard can be confused 

with ground ivy leaves when the plants are not in flower (Fig. 

61). Look closely for hair on the leaves. Ground ivy leaves are 

hairy whereas garlic mustard leaves are not. These plants 

cannot be confused when they are in flower because ground ivy 

has purple flowers (Fig. 62) whereas garlic mustard has white 

flowers (Newcomb, 1977; Dickinson et al. 2004). 

Figure 61: Comparing the leaves of ground 

ivy 1 (left) to those of garlic mustard 1 (right). 

violets (Viola spp.) 

Figure 62: Ground ivy flowers 1 . 

Violet leaves can be confused with the first-year basal leaves of 

garlic mustard when the plants are not in flower (Fig. 63). It is 

often very difficult to differentiate between violets and garlic 

mustard. Some guides point out that the wavy edges are less 

than 1mm long for violets whereas they are greater than 1mm 

long for garlic mustard. Perform a quick check by crushing 

the leaves to detect whether or not they emit a garlic odour. 

Although some violet species have white flowers they are not 

similar to those of garlic mustard (Newcomb, 1977; Dickinson 

et al. 2004) (Fig. 64). 

Figure 63: Comparing violet leaves 1 (top) 

to garlic mustard basal leaves 1 (bottom). 

Figure 64: Violet flowers 20 .


kidney-leaved buttercup (Ranunculus abortivus) 

Before flowering, the first-year basal leaves of garlic mustard 

can be confused with the basal leaves of kidney-leaved 

buttercup (Fig. 65). The leaves of kidney-leaved buttercup have a 

very flat appearance whereas the basal leaves of garlic mustard 

have deep veins that give them a wrinkled appearance. Kidneyleaved 

buttercup flowers are yellow (Fig. 66) in contrast to the 

white flowers of garlic mustard (Newcomb, 1977; Dickinson et 

al. 2004). 

Figure 65: Comparing the basal leaves of 

kidney-leaved buttercup 1 (left) to those of garlic 

mustard 1 (right). 

Figure 67: Comparing the leaves 

of wild ginger 1 (top) to those of 

garlic mustard 1 (bottom). 

wild ginger (Asarum canadense) 

Figure 66: Kidney-leaved buttercup flower 1 . 

Wild ginger leaves may appear to be similar to garlic mustard 

basal leaves at first sight (Fig. 67). However, wild ginger leaves 

do not have the wavy edges that are characteristic of the basal 

leaves of garlic mustard. The flowers of wild ginger emerge 

close to the ground and are red (Fig. 68). Garlic mustard flowers 

are white and grow on the top of flowering stalks (Newcomb, 

1977; Dickinson et al. 2004). 

Figure 68: Wild ginger flower 1 . 

[ 73 ]

[ 74 ] 


toothwort (Cardamine spp.) 

The flowers of toothwort are very similar in appearance to garlic 

mustard flowers. Both species have white flowers with four 

petals (Fig. 69). In contrast, these plants are easily identified 

by the characteristics of their leaves. Toothwort leaves are 

compound, consisting of three leaflets (Fig. 70), whereas garlic 

mustard leaves are simple (Newcomb, 1977; Dickinson et al. 

2004). 

Figure 69: Comparing the flowers of toothwort 1 (left) to those of garlic mustard 13 (right). 

Figure 70: Comparing toothwort 1 (left) and garlic mustard 1 (right). 

catnip (Nepeta cataria) 

Catnip leaves may be confused with the second-year leaves 

of garlic mustard. They are both triangular with sharp edges 

(Fig. 71). Close inspection will reveal that catnip leaves are very 

hairy while garlic mustard leaves are not. Catnip flowers do not 

resemble garlic mustard flowers (Fig. 72) as they are irregular 

in shape (Newcomb, 1977; Dickinson et al. 2004). 

Figure 71: Comparing the leaves of catnip 1 (left) to the second-year leaves of 

garlic mustard 1 (right). 

Figure 72: Catnip flowers 1


A key to plants that may be confused with garlic 

mustard (Alliaria petiolata) 

1..Compound.leaves......................................................................................... Toothwort.(Cardamine spp.) 

1..Simple.leaves. 

. 2..Leaf.edges.smooth................................................................................... Wild.ginger.(Asarum canadense) 

. 2..Leaf.edges.toothed 

. . 3..Teeth.rounded 

. . . 4..Leaves.with.hair................................................................................ Ground.ivy.(Glechoma hederacea) 

. . . 4..Leaves.without.hair 

. . . . 5..Deep.veins.give.leaves.a.wrinkled.appearance................ Garlic.mustard.(Alliaria petiolata) 

. . . . 5..Leaves.appear.flat....................................................................... Kidney-leaved.buttercup.(Ranunculus abortivus) 

. . 3..Teeth.sharp.or.pointed 

. . . . . 6..Leaves.very.hairy.................................................................... Catnip.(Nepeta cataria) 

. . . . . 6..Leaves.with.little.or.no.hair 

. . . . . . 7..Teeth.less.than.1mm.long............................................... Violet.(Viola spp.) 

. . . . . . 7..Teeth.greater.than.1mm.long........................................ Garlic.mustard.(Alliaria petiolata) 


Kingdom. Plantae 

...Subkingdom. Tracheobionta 

......Division. Magnoliophyta 

........Class. Magnoliopsida 

.........Subclass. Dilleniidae 

...........Order. Brassicales 

............Family. Brassicaceae 

.............Genus. Alliaria 

...............Species. Alliaria petiolata 

BioloGy 


A native to Eurasia, garlic mustard 

was probably introduced to North 

America by early settlers. It was 

valued for its garlic flavour and 

healing properties (Huffman, 2005). 

The first documented evidence of 

garlic mustard in North America 

was found on Long Island, New 

York in 1868 (Anderson et al. 1996). 

Canadian records show that garlic 

mustard was found in Toronto, 

Ontario in 1879 where it has since 

spread to Quebec, New Brunswick, 

Nova Scotia and British Columbia 

(Cavers et al. 1979). 

[ 75 ]

[ 76 ] 


hABitAt 

Garlic mustard is a shade tolerant species (Rodgers et al. 

2008) that grows best in moist soils (Cavers et al. 1979). In its 

native range, garlic mustard occupies edge habitats such as 

those found on the borders of forests and rivers (Rodgers et 

al. 2008). In North America, garlic mustard also prefers these 

edge habitats. However, populations can be found anywhere 

from forest interiors to open fields (Cavers et al. 1979) (Fig. 73). 

Disturbed areas allow seedlings to establish where they can 

spread and invade undisturbed habitats (Kaufman & Kaufman, 

2007). 

Figure 73: Garlic mustard in the forest understory 1 . 

reProduction 

Garlic mustard plants reproduce via seed production. Although 

a variety of insects help to pollinate the flowers of garlic 

mustard, these plants can also self-pollinate. This ensures that 

seeds will be produced by every flowering individual (Rodgers 

et al. 2008). Mature plants produce seedpods that hold 10 to 20 

seeds and each plant can have anywhere from 1 to 150 seedpods 

(Cavers et al. 1979). Seeds have a thick outer layer that allows 

them to persist in the soil for up to 10 years (Rodgers et al. 

2008). 

liFe cycle 

Garlic mustard has a biennial (two-year) life cycle. Basal leaves 

develop during the first year. Flowers emerge in April during 

the second year of growth. Seedpods are formed shortly 

thereafter and seeds are dispersed throughout the summer 

and fall (Cavers et al. 1979).



Several aspects relating to garlic mustard’s reproductive 

capabilities lend to its success as an invader. These include the 

ability to self-pollinate, to produce a large quantity of viable 

seed, and to form extensive seed banks. Many native plants 

need insect pollinators in order to produce seed. However, 

garlic mustard has the ability to self-pollinate. This gives 

garlic mustard the advantage of consistent seed production 

(Rodgers et al. 2008). Populations of garlic mustard can 

increase rapidly because these plants have a high reproductive 

potential (Drayton & Primack, 1999). An individual plant has 

the ability to produce over 3500 seeds and studies have shown 

that several populations in Ontario can produce 107 000 seeds 

within one square metre (Cavers et al. 1979). After dispersal, 

seeds remain dormant until ideal conditions prompt them to 

germinate. A thick outer seed coat (Cavers et al. 1979) allows 

seeds to remain viable for up to 10 years, forming a seed bank 

in the soil (Rodgers et al. 2008). 

Garlic mustard has the ability to thrive in areas with a wide 

range of different light intensities. The basal leaves of garlic 

mustard remain green all year providing the plant with 

potential photosynthetic activity during late fall and early 

spring when other native herbs are dormant (Myers et al. 2005) 

(Fig. 74). Photosynthesis is important because it allows plants 

to convert solar energy into organic compounds for growth 

and reproduction. Being an evergreen species, garlic mustard 

obtains a competitive advantage over other species that can 

only photosynthesize during the summer months (Myers et al. 

2005). 

Figure 74: Garlic mustard’s basal leaves remain green all year long 1 . 

[ 77 ]

[ 78 ] 


Garlic mustard has a variety of secondary compounds that may 

decrease the likelihood of herbivore attack and suppress the 

growth of competing plants. Secondary compounds are found 

in the plant tissues and can react with water to form cyanide 

compounds. These compounds are toxic and help to discourage 

potential herbivores from feeding on the plants (Rodgers et al. 

2008). In its native range, organisms have had time to co-evolve 

with these toxic compounds whereas North American species 

have not (Renwick, 2002). 

Secondary compounds can potentially suppress the growth of 

competing native plants. This may be accomplished indirectly, 

by decreasing the amount of mycorrhizal fungi found in the 

soil (Barto et al. 2010). Unlike most native plants that rely on 

these beneficial fungi for nutrients, garlic mustard does not 

establish associations with mycorrhizal fungi. As such, by 

depressing mycorrhizal associations of native plants they can 

gain a competitive advantage (Wolfe et al. 2008). 

ecoloGicAl iMPActs 

Garlic mustard may change the species composition in mature 

hardwood forests. Greenhouse studies have shown that garlic 

mustard can dramatically affect the growth rate of dominant 

hardwood species such as sugar maple (Acer saccharum), 

red maple (Acer rubrum) and white ash (Fraxinus americana). 

Regeneration of these canopy tree species is repressed which 

in time could change the overall composition of a mature 

hardwood stand to that of an early successional community 

(Stinson et al. 2006). Further research in the field is needed to 

assess the effects of garlic mustard on the species composition 

of mature forests. 

Diversity within a forest stand is important because it supports 

a healthy ecosystem. A diverse forest is less susceptible to the 

impacts of pest outbreaks and disease (Chapeskie et al. 2006). 

Garlic mustard actively decreases diversity through its superior 

competitive abilities and by preventing nutrient uptake by 

mycorrhiza-dependent species (Stinson et al. 2006). Dense 

stands can form in the understory, effectively decreasing the 

abundance of native wildflowers and potentially repressing 

hardwood regeneration (Fig. 75). 

Garlic mustard invasions affect wildlife. Studies have shown 

that garlic mustard has a negative impact on certain species 

of butterfly. Native species such as West Virginia white (Pieris 

virginiensis) and mustard white (Pieris napi oleracea) have 

been observed to deposit their eggs on garlic mustard plants. 

Unfortunately, secondary compounds greatly lower the survival 

rate of hatching larvae (Rodgers et al. 2008). These butterflies


usually oviposit on native toothwort (Cardamine sp.) found in 

forest understories. Garlic mustard populations can encroach 

upon toothwort habitat and gradually displace toothwort 

populations. With the absence of toothwort, these butterflies 

have little choice but to deposit their eggs on garlic mustard 

(Renwick, 2002). 

Figure 75: Second-year garlic mustard plants towering over native 

vegetation 1 . 

vectors/PAthwAys 

Garlic mustard seeds usually fall within several metres of the 

parent plant. This allows for the growth of dense patches and the 

development of a large seed bank for established populations 

(Anderson et al. 1996). As populations have been observed 

along flood water paths it seems likely that flood waters can 

increase dispersal distance (Wilkens, 2000). 

Long distance dispersal usually occurs with the help of other 

organisms. Humans are mainly responsible for the dispersal 

of garlic mustard seeds. Seeds can readily attach to clothing, 

shoes and tires. Seeds can get stuck to machinery such as heavy 

equipment used for creating roads or used for logging. Tires 

from all-terrain vehicles, tractors and trailers contribute to 

seed dispersal (Shartell et al. 2011). 

Animals such as mice, raccoons and squirrels have been 

observed collecting seeds and moving them to roosting areas. 

Seeds often get stuck to the hooves of deer where they can be 

carried over long distances (Wilkens, 2000). While feeding, deer 

will often create areas of disturbance where garlic mustard can 

easily become established (Rodgers et al. 2008). 

[ 79 ]

[ 80 ] 




• Prevent seeds from entering the woodlot by limiting access and 

washing all shoes and tires before entry; 

• Do not let domestic animals roam freely if there are known garlic 

mustard populations in or around the woodlot; 

• Try to minimize soil disturbance during woodlot management 

activities; 

• Volunteer to help eliminate garlic mustard populations from 

nearby properties. 


• Monitor the woodlot frequently paying attention to roadsides, 

flood water paths and disturbed areas; 

• Learn how to identify both the basal rosette stage and secondyear 

mature stage of garlic mustard. 


Hand-pulling: Garlic mustard can be pulled from the ground 

with relative ease. Pulling from the base of the stalk will 

prevent the stem from breaking and allow a large majority of 

the root to be removed. Remove as much of the root as possible, 

with the upper half of the taproot being removed at the very 

least, to prevent re-sprouting (Fig. 76). All above-ground plant 

tissue should be placed in a bag and removed from the area. 

This is essential as pulled stems may have the ability to release 

viable seed. Care should be taken not to trample or pull native 

vegetation (NCC, 2007). 

Figure 76: Hand-pulling garlic mustard plants 1 .


In areas with small invasions or scattered satellite populations, 

the goal of hand-pulling should be to remove both first and 

second-year plants. In areas with larger invasions it will 

become impractical to try to remove all garlic mustard plants. 

In these areas, focus should lean towards removing all mature 

second-year plants while allowing the basal rosettes to remain. 

Removing the second-year plants will prevent seeds from 

producing and replenishing the seed bank. Management will 

need to continue for years until the seed bank is completely 

exhausted (Nuzzo, 1991). 

Place all plant parts in a plastic or paper bag and dispose of in 

an appropriate landfill (Fig. 77). Garlic mustard is not a yard 

waste and should not be disposed of as such. Contact your 

local waste service provider and/or landfill manager to discuss 

disposal procedures. Ensure that no parts are left behind to 

produce roots or seed. Make sure that bags are properly sealed 

to prevent spread during transportation (Frey et al. 2005). 

Figure 77: Garlic mustard disposal 1 . 

Flower-head removal: In small populations, removing flower 

heads can prevent seed production. However, constant 

vigilance is required as clipping encourages the growth of 

more flowers. This control method should only be used in 

very small populations as it involves frequent monitoring. 

One advantage of removing flower heads over hand-pulling is 

the absence of soil disturbance. When plants are pulled, they 

create extra space on the forest floor where garlic mustard can 

reinvade (NCC, 2007). Care should be taken when selecting 

this management option. It may prove difficult to ensure that 

all flowers have been removed. Missing a single reproductive 

individual could potentially negate the time and effort invested 

in control if seeds are allowed to replenish the seed bank (Frey 

et al. 2005). 

Mowing or cutting: Manual control of dense patches can be 

accomplished by mowing or cutting. This is only appropriate in 

areas where large invasions have eliminated native vegetation. 

[ 81 ]

[ 82 ] 


Cutting can be done using weed whackers, scythes, and lawn 

mowers. The objective is to cut as close to the ground as 

possible. This control method will need to be conducted several 

times throughout the season as new shoots arise from the 

taproots (Drayton & Primack, 1999). There is a short window 

of opportunity in which the most damage can be done to 

individual garlic mustard plants using the cutting method. The 

roots of garlic mustard allocate the greatest amount of energy 

to the flower and fruiting stage, thus cutting the plants right 

after they produce flowers may leave the remaining roots in 

an energy deficient state (NCC, 2007). Cutting should be done 

before seeds are produced and throughout the season to prevent 

any late seedpod production. The main goal is to exhaust the 

seed bank, gradually eliminating the entire population (Strobl 

& Bland, 2000). 

Herbicide application: Dense patches can be controlled 

chemically. Studies have shown that herbicides can decrease 

garlic mustard populations by as much as 91% (Rodgers et 

al. 2008). In order to decrease the amount of damage done to 

native vegetation it would be appropriate to use a spot-treatment 

method. This may increase the amount of labour required, 

however, it is essential to minimize any impacts to native 

plants (Landis & Evans, 2009). Herbicide application should 

only be done in the spring or fall when most native plants are 

dormant. A spring application has the advantage of eradicating 

both the first-year basal rosettes and the second-year flowering 

plants (Slaughter et al. 2007). However, a spring application may 

be inappropriate if the affected woodlot is valued for its early 

spring ephemerals (NCC, 2007). A fall application is generally 

preferred because a large majority of native plants have already 

entered into dormancy (Slaughter et al. 2007). 




A plan should be created before the commencement of any 

management practice. Control efforts should focus on working 

from the outer boundaries of the invasion towards the centre. 

This will prevent seeds, a main dispersal agent for garlic 

mustard, from being spread to other areas of the hardwood 

stand. Be sure to inspect and clean footwear, clothing, tires and 

any other objects that could harbour seeds before moving to 

other areas (Nuzzo, 1991). 

In the spring it is best to hand-pull, cut or mow dense patches of 

second-year plants shortly after they begin to flower but before


they begin producing seedpods. Use caution in areas where 

there is desirable vegetation to prevent any unwanted damage. 

The goal is to prevent any seeds from replenishing the seed 

bank. Monitor the area throughout the summer and pull any late 

flowering individuals (NCC, 2007). Apply herbicides in the fall, 

after the majority of native vegetation has entered dormancy. 

Targeting the basal rosettes should reduce the number of 

second-year plants produced the following year. Repeat for 2 to 

3 years or until the population has reached a manageable size 

to fully rely upon non-chemical control methods such as handpulling 

(Slaughter et al. 2007). 


Hand-pull, cut or mow dense patches of second-year plants 

shortly after they begin to flower in the spring. Ensure this 

is done before the garlic mustard plants begin producing 

seedpods. Use caution in areas where there is desirable 

vegetation to prevent any unwanted damage. The goal is to 

prevent any seeds from replenishing the seed bank. Monitor 

the area and pull any remaining second-year plants throughout 

the summer. Focus on eliminating second-year plants to slowly 

exhaust the seed bank. As the population begins to decrease 

in size, basal rosettes may also be targeted to help speed up 

control (Nuzzo, 1991). 

There is the potential to reintroduce native species to these 

disturbed areas which may jumpstart the restoration of the 

forest ecosystem. Results from a study by Murphy (2005) revealed 

that planting a native wildflower, bloodroot (Sanguinaria 

canadensis), helped to alleviate the spread of garlic mustard 

(Fig. 78). Further research is needed to determine which native 

species would be appropriate for such restoration efforts. 

Figure 78: Planting a native species, bloodroot (Sanguinaria canadensis) 1 . 

[ 83 ]

[ 84 ] 


Table 8: Management recommendations for garlic mustard (Alliaria petiolata). 

Extent.of.infestation Small.invasions.and.satellite.populations Large.invasions.and.dense.populations 

Recommended. 


Hand-pulling.and.flower.head.removal. Integrated.control.including.physical. 

control.methods.and.herbicide.. 

application. 

Timing Hand-pulling:.Spring.(April-May). 

Flower.head.removal:.Spring.and.early. 

summer.(April-June). 

Frequency.of.control Hand-pulling:.Several.times.per.year,.up.to. 

five.years.or.until.the.seed.bank.is.exhausted. 

Flower.head.removal:.Every.few.days.until.no. 

more.flowers.are.produced. 

Length.of.control 2-5.years. 5.+.years. 

required.restoration Plant.native.species.in.areas.where.handpulling.creates.soil.disturbance. 

Integrated.control:.. 

Physical.control:.Spring.and.summer.. 

Herbicide.application:.Fall. 

Manual.removal.in.the.spring.and.chemical. 

control.in.the.fall.may.be.required.for.up.to. 

5.years.or.until.seed.bank.is.exhausted. 

Plant.native.species.once.seed.bank.has. 

nearly.been.exhausted..Monitoring.and. 

hand-pulling.any.germinating.garlic. 

mustard.seedlings.will.be.needed.


5.1.4 

Barberry 

(Berberis thunbergii & B. vulgaris) 


japanese barberry, Purple japanese barberry, common barberry, 

european barberry, jaundice berry 


Figure 79: Japanese barberry 1 . 

85 

[ ]

[ 86 ] 



Two species of barberry currently invade Ontario hardwood 

stands: Japanese barberry (Berberis thunbergii) and common 

barberry (Berberis vulgaris). They are both deciduous shrubs 

with yellowish bark and spiny branches (Bailey, 1969). 

japanese barberry (Berberis thunbergii) 

steM & BrAnches: The branches are brown with deep 

grooves and have alternately arranged simple thorns (Fig. 80). 

Japanese barberry shrubs range from 0.5m to 2m in height 

(Bailey, 1969). 

Figure 80: Japanese barberry stem with simple thorns 1 . 

leAves: Japanese barberry has simple, oval leaves with 

smooth margins which taper at the base (Fig. 81). The leaves are 

found in clusters of approximately 2 to 6 leaves located above 

each thorn. Leaves range in colour from green to red to purple, 

depending on the cultivar (Kaufman & Kaufman, 2007). 

Figure 81: Japanese barberry leaves 1 .

5.1.4 BARBERRY (Berberis thunbergii & B. vulgaris) 

Flowers: Pale yellow flowers emerge from the leaf axils, 

either as a solitary flower or in groups of 2 to 4 (Swearingen et 

al. 2010). They have 6 petals, 6 sepals and 6 stamen (Fernald, 

1950) (Fig. 82). 

Figure 82: Japanese barberry flowers 1 . 

Fruit/seed: The fruit is a bright red, oval berry containing 

several small seeds (Bailey, 1969) (Fig. 83). 

common barberry 

(Berberis vulgaris) 

Figure 83: Fruit of Japanese barberry 1 . 

steMs & BrAnches: The branches 

of common barberry are brownish-grey 

with deep grooves and have alternately 

arranged 3-pronged spines (Bailey, 1969) 

(Fig. 84). Common barberry shrubs range 

from 1 to 3m in height (Fernald, 1950). 

Figure 84: Common barberry stem 

with 3-pronged spines 6 . 

[ 87 ]

[ 88 ] 


Figure 85: Common 

barberry leaves 6 . 

leAves: Common barberry has simple, oval leaves with 

toothed margins which taper at the base (Fig. 85). They are 

found in clusters along the stem (Bailey, 1969). Leaves are green 

but turn red, orange or purple in the fall (Gucker, 2009). 

Flowers: Clusters of 10 to 20 pale yellow flowers grow in 

racemes (Gleason & Cronquist, 1963). They have 6 petals, 6 

sepals and 6 stamen (Fernald, 1950) (Fig. 86). 

Fruit/seed: The fruit is a berry containing several small 

seeds (Bailey, 1969). As the fruits mature they change from 

green to bright red (Fig. 87). 

Figure 86: Common barberry 

flower clusters 6 . 


Barberries are easily distinguished from other shrubs during 

the growing season when they have leaves. The oval, tapering 

leaves found in clusters along the stem are different than those 

of native shrubs in Ontario. However, the thorny branches may 

be confused with those of other shrubs when the leaves are 

absent during the colder months. Similar shrubs with thorns 

include buckthorn (Rhamnus spp.) hawthorn (Crataegus spp.), 

and gooseberry (Ribes spp.). 

Buckthorn (Rhamnus spp.) 

Several species of native and exotic buckthorn are found in 

Ontario. Buckthorn can be easily distinguished from barberry 

by the position of its thorns, which are opposite to one another 

while barberry has alternately arranged thorns (Fig. 88). The 

leaves of buckthorn are different than those of barberry; they 

have an opposite arrangement and do not taper at the base 

(Symonds, 1963) (Fig. 89). 

Figure 87: Common 

barberry fruit 14 .


Figure 88: Comparing the opposite thorns of buckthorn 14 (left) to the alternately arranged thorns of 

Japanese barberry 1 (right). 

Figure 89: Comparing a buckthorn leaf 1 (left) to that of Japanese barberry 1 (right). 

hawthorn (Crataegus spp.) 

There are many species of native hawthorn in Ontario. These 

hawthorns can sometimes be confused with barberries because 

they also have alternately arranged single thorns. However, their 

toothed leaves contrast with those of Japanese barberry, which 

have smooth edges (Fig. 90). Common barberry has 3-pronged 

thorns whereas hawthorns have singular thorns (Fig. 91). Unlike 

barberries, the buds of hawthorn are almost spherical and are 

not located above each thorn (Symonds, 1963). 

Figure 90: Comparing the thorns and leaves of 

hawthorn 21 (top) to those of Japanese barberry 1 (bottom). 

Figure 91: Comparing the singular thorns of hawthorn 16 

(left) to the 3-pronged thorns of common barberry 6 (right). 

[ 89 ]

[ 90 ] 


A key to shrubs that may be confused 

with barberry (Berberis spp.) 

1..Leaves.are.present.(spring,.summer,.fall) 

. 2..Leaves.with.toothed.edges................................................................................Common.barberry.(Berberis vulgaris) 

. 2..Leaves.with.smooth.edges.................................................................................Japanese.barberry.(Berberis thunbergii) 

1..Leaves.are.not.present.(winter) 

. . 3..Thorns.found.opposite.to.one.another.....................................................Buckthorn.(Rhamnus.spp.) 

. . 3..Thorns.with.an.alternate.arrangement 

. . . 4..Buds.almost.spherical.................................................................................Hawthorn.(Crataegus.spp.) 

. . . 4..Buds.not.spherical 

. . . . 5..Unbranched,.single.spine......................................................................Japanese.barberry.(Berberis thunbergii) 

. . . . 5..Branched,.3-pronged.spines................................................................Common.barberry.(Berberis vulgaris) 


Japanese barberry.(Berberis thunbergii) 




......Class. Magnoliopsida 

........Subclass. Magnoliidae 

..........Order. Ranunculales. 

............Family. Berberidaceae 

..............Genus. Berberis 

................Species. Berberis thunbergii 

BioloGy 


Common barberry.(Berberis vulgaris) 





........Subclass. Magnoliidae. 

..........Order. Ranunculales 

............Family. Berberidaceae 

..............Genus. Berberis 

................Species. Berberis vulgaris 

Common barberry was introduced by early settlers due to its 

popular uses in dyes and jams. However, when it was discovered 

to act as a host for cereal stem rust (Puccinia graminis), control 

measures were put in place to eradicate the shrub from North


America (Kaufman & Kaufman, 2007). An eradication program 

was initiated in 1964 in Ontario and Quebec but complete 

eradication was never achieved. Since the 1980’s populations 

have begun to increase due to alleviation of control measures 

(Clark et al. 1986). 

Japanese barberry was brought into the country from Japan as 

an alternative to common barberry in the late 1800’s. Although 

it is not a host for any agricultural crop diseases it has invaded 

forest understories. In spite of this outcome, some cultivars of 

Japanese barberry are still being sold as ornamental shrubs in 

Ontario (Zouhar, 2008). 

hABitAt 

Japanese barberry has high phenotypic plasticity, growing in 

a wide array of light and moisture conditions (Silander Jr. & 

Klepeis, 1999). It has been documented in riparian areas and 

in both open and closed-canopy habitats such as wetlands, old 

fields and forests (Zouhar, 2008). Japanese barberry can escape 

from cultivation and form dense stands in undisturbed forest 

understories (Ehrenfeld, 1997) (Fig. 92). 

Figure 92: Japanese barberry growing in the forest understory 6 . 

reProduction 

Barberry can spread via sexual and vegetative reproduction. 

Barberries produce a multitude of fruit. Seeds are dispersed as 

berries drop to the ground or are eaten by wildlife. Barberries 

reproduce vegetatively by way of tillers produced from the root 

system or when stems take root when they come into contact 

with the ground (Ehrenfeld, 1999). 

[ 91 ]

[ 92 ] 


liFe cycle 

Barberry seeds germinate in May. Mature shrubs produce 

flowers from April to June. Berries mature in late summer, 

usually at the end of August and early September. Unless eaten 

by wildlife, these berries will remain on the plants throughout 

the winter (Ehrenfeld, 1999) (Fig. 93). 

Figure 93: Japanese barberry fruits are often seen in winter 14 . 


Barberry has high recruitment rates due to its ability to 

produce large quantities of seed. As stem densities increase in 

a barberry stand so does the rate of recruitment. The ability to 

employ a variety of reproductive methods (see above) increases 

barberry’s ability to invade an area. Populations can expand 

quickly, taking up space and shading-out native understory 

species (Ehrenfeld, 1999). 

Barberry produces leaves very early in the season before other 

native shrub species and well before canopy closure (Fig. 94). 

This results in longer photosynthetic activity compared to 

other co-occurring species (Silander Jr. & Klepeis, 1999) and 

helps these invasive shrubs to become established in the forest 

understory before light levels become sub-optimal as the 

canopy closes (Cheng-Yuan et al. 2007). 

Figure 94: Japanese barberry in early spring 6 .



Barberry forms dense 

stands in hardwood forest 

understories (Fig. 95). Such 

stands cover large areas and 

compete with native plants 

for space, nutrients and 

light (Zouhar, 2008). One 

study revealed a positive 

correlation between Japanese 

barberry stands and blacklegged 

tick (Ixodes scapularis) 

populations. Indeed, areas 

invaded by Japanese barberry 

supported a greater number 

of tick hosts (Elias et al. 

2006). Black-legged ticks are 

detrimental to human health 

as they can transmit a variety 

of diseases such as Lyme 

disease (Borrelia burgdorferi) 

(Magnarelli et al. 2006). 

Barberry populations can change forest soil characteristics 

by increasing pH levels and nitrification rates (Ehrenfeld et 

al. 2001). These edaphic effects may inhibit the restoration of 

native flora and alter the forest’s successional patterns after 

the removal of this invasive plant (Kourtev et al. 1999; Zouhar, 

2008). 


Figure 95: Japanese barberry invasion 22 . 

Barberry has bright red berries, which are attractive to wildlife. 

Turkeys, grouse and various songbirds consume the berries. 

Berries may thus be transported to other places where the birds 

eat the fruit pulp and discard the seeds. Seeds may also be eaten 

and passed through the digestive tract to later be discarded, 

often quite far from their origin (Silander Jr. & Klepeis, 1999). 

Animals such as chipmunks, mice and deer may also play a 

role in seed dispersal (Ehrenfeld, 1997; 1999). 

Common barberry cannot be imported or sold within Canada 

due to its ability to act as an alternate host for cereal stem rust 

disease (Puccinia graminis) (Fig. 96). However, twelve cultivars 

of Japanese barberry have been approved for sale in Canada 

because they have been found to be resistant to the disease. 

Sold as ornamental shrubs, these cultivars frequently escape 

from cultivation due to bird-mediated dispersal. Humans can 

[ 93 ]

[ 94 ] 


aid in seed dispersal when seeds get stuck under shoes or in 

tire treads (CFIA, 2008). 

Figure 96: The symptoms of cereal stem rust (Puccinia graminis) 23 . 



• Barberry is considered an invasive plant and should not be planted 

in the garden. Use native alternatives or species that are not 

considered invasive; 


washing shoes and tires before entry. 


• Learn how to identify both Japanese and common barberry; 

• Regularly monitor the woodlot for new invasions and pay particular 

attention to road sides and trails where invasive plants can first 

establish; 

• Talk with neighbours about what they grow in their gardens and the 

threats associated with growing invasive plants. 


Hand-pulling, excavation and cutting: Although hand-pulling 

is a common method for controlling small populations of 

invasive plants, hand-pulling barberry shrubs is impractical 

due to their large size. Excavation with root wrenches allows


roots to be dug up from the soil which can be effective but time 

consuming and labour intensive, unless the population is very 

small. Ensure that all stem fragments are collected as shoots 

can easily re-sprout. Cutting is suggested in areas where dense 

populations of barberry create access problems. Other methods 

of control will need to be administered to the exposed stumps 

to prevent re-sprouting (Silander Jr. & Klepeis, 1999; Ward et al. 

2009). 

Directed flame treatment: Applying a directed flame to the 

base of the stump is effective and minimizes disturbance, 

especially when compared with excavation. Using a propane 

torch and applying flame for 20 seconds to the base of the 

stem can effectively kill small shrubs in a single application 

(Ward et al. 2009). However, larger shrubs may require followup 

treatments. The directed flame method should only be 

attempted when the forest floor is wet or damp to prevent the 

risk of a fire (Ward et al. 2010). Consult with your local fire 

department before using the directed flame method. 

Herbicide application: Herbicides may be applied as a foliar 

spray or using the basal bark or cut-stump methods. The 

latter methods are preferable as they reduce herbicide drift to 

nearby native vegetation. Basal bark treatments are effective 

and generally less labour intensive than the cut-stump method. 

Ensure that chemical treatments are performed in both the 

spring and fall for effective control of barberry invasions 

(Silander Jr. & Klepeis, 1999). 




Control of barberry populations should occur twice per year, 

once in the spring and once during the fall. The goal of control 

treatments in the spring is to remove as many barberry shrubs 

as possible, paying particular attention to large, mature plants. 

Control treatments in the fall can focus on newly germinating 

seedlings as well as on any individuals that were not removed 

in the spring. 

Herbicides can be applied in the spring. For large invasions, 

apply herbicides as a foliar spray. Barberry leaves emerge earlier 

in the spring than those of co-occurring native plants. This is 

an ideal time to apply herbicides because there is a lower chance 

of non-target effects on native vegetation. Herbicide application 

may prove difficult in areas where barberry populations exist 

as dense shrub thickets. In this case, it may be easier to first cut 

the stems close to the ground which would allow better access 

to the area. Afterwards, herbicides can be applied to the cut 

stems. In the fall, control will consist of treating any remaining 

[ 95 ]

[ 96 ] 


individuals, newly germinating seedlings and re-sprouting 

shoots. Basal bark treatments are effective in areas where 

populations are not so dense that access is achievable. Basal 

bark treatments can be more cost effective as less herbicide 

is required and there is little chance of incidental damage to 

other plants (Silander Jr. & Klepeis, 1999). 


If chemical control is not possible or wanted, the directed flame 

method is often a successful means of control for barberry 

invasion. In dense patches where access is difficult, cutting 

first is suggested. This is followed by removal of the woody 

stems, thereby allowing the base of the stump to be exposed 

for application of a directed flame. In areas where access is 

not a problem, stems do not need to be cut as the flame can 

be directed to the base. Directed flame treatments in both the 

spring and fall can be a cost-effective and relatively simple 

means of controlling barberry populations. The directed flame 

method should only be used when the forest floor is wet or 

damp to prevent the risk of a fire (Ward et al. 2010). Remember 

to consult with the local fire department before using the 

directed flame method. 

Table 9: Management recommendations for barberry (Berberis spp.). 

Extent.of.infestation Small.invasions.and.satellite.populations Large.invasions.and.dense.populations. 

Recommended. 


Excavation.or.directed.flame. Cutting.followed.by.directed.flame. 

and/or.chemical.control. 

Timing Spring.and.fall. Spring.and.fall. 

Disposal Remove.woody.debris.to.a.brush.pile.for. 

burning..Dead.standing.shrubs.may.be. 

left.on.site.or.cut.and.removed. 

Frequency.of.control Twice.yearly. Twice.yearly. 

Length.of.control 1-2.years. 1-3.years. 

Required.restoration Plant.native.species.in.areas.where. 

excavation.creates.soil.disturbance. 

Remove.woody.debris.to.a.brush.pile. 

for.burning..Dead.standing.shrubs.may. 

be.left.on.site.or.cut.and.removed. 

Active.seeding.or.planting.with.native. 

species.may.be.needed.


5.1.5 

japanese knotweed 

(Fallopia japonica) 


japanese fleece flower, Mexican bamboo, crimson beauty, reynoutria 


Figure 97: Japanese knotweed 1 . 

97 

[ ]

[ 98 ] 



steM: Japanese knotweed has large, bamboo-like, hollow 

stems with green, red and purple markings. Distinctive nodes 

can be seen along the entire stem (Weber, 2003) (Fig. 98). 

Figure 98: Japanese knotweed stems 1 . 

leAves: Japanese knotweed has simple triangular leaves 

arranged alternately along the stem (Wilson, 2007) (Fig. 99). 

Figure 99: Japanese knotweed leaves 1 . 

Flowers: Japanese knotweed produces numerous flowers 

arranged in clusters. Each flower has 5 white petals (Fig. 100). 

Flowers emerge in late summer and persist through the fall 

(Bailey, 1969). 

Figure 100: Japanese knotweed flowers 1 .

5.1.5 JAPANESE KNOTWEED (Fallopia japonica) 

Fruit/seed: The fruits are 3-sided and winged, encasing 

a single brown seed (Weber, 2003) (Fig. 101). There have been 

conflicting results regarding the viability of Japanese knotweed 

seeds. As such, more research is needed to determine the extent 

to which the seeds produced by this invasive species contribute 

to its spread. 

roots: Japanese knotweed produces a very large and extensive 

rhizome from which multiple stems arise (Fig. 102). This creates 

dense patches that consist of a single individual (Bailey, 1969). 

heiGht: Japanese knotweed is a tall growing plant that can 

often reach 4m in height (Wilson, 2007) (Fig.103). 

Figure 101: The fruit of Japanese knotweed 4 . Figure 102: Japanese knotweed roots 19 . 

Figure 103: Japanese knotweed can be up to 4m tall 1 . 

[ 99 ]

[ 100] 


dock (Rumex spp.) 


The stems of some Rumex species may look very similar to 

those of Japanese knotweed. Species such as great water dock 

(R. orbiculatus) have jointed stems that can grow up to 2m tall 

and may even appear red at times (Fig. 104). Both great water 

dock and Japanese knotweed have fruits that are 3-sided 

(Fig. 105). However, Rumex species generally have long, lanceshaped 

leaves whereas Japanese knotweed has triangular leaves 

(Newcomb, 1977). 

Figure 104: Comparing the stems of dock 20 (left) to those of Japanese knotweed 4 (right). 

Figure 105: Comparing the seeds of dock 20 (left) to those of Japanese knotweed 6 (right). 

Giant hogweed (Heracleum mantegazzianum) 

Giant hogweed is an introduced species from Eurasia. Like 

Japanese knotweed, its hollow stems are green with small 

purple spots and can reach approximately 5m in height (Fig. 

106). In steep contrast, giant hogweed has deeply lobed leaves


whereas those of Japanese knotweed are triangular and 

unlobed (Kaufman & Kaufman, 2007) (Fig. 107). 

Figure 106: Comparing the stems of giant hogweed 6 

(left) to those of Japanese knotweed 1 (right). 

common cow parsnip (Heracleum maximum) 

Figure 107: Comparing the leaves of giant hogweed 24 

(top) to those of Japanese knotweed 1 (bottom). 

Common cow parsnip belongs to the same genus as giant 

hogweed. However, it is native to North America. As with both 

giant hogweed and Japanese knotweed, common cow parsnip 

has tall, hollow stems. Common cow parsnip has deeply lobed 

leaves in contrast to the triangular leaves of Japanese knotweed 

(Newcomb, 1977) (Fig. 108). 

Figure 108: Comparing common cowparsnip 1 (left) and Japanese knotweed 1 (right). 

[ 101]

[ 102] 


wild buckwheat (Fallopia convolvulus) 

Wild buckwheat and Japanese knotweed have similar triangular 

leaves. However, wild buckwheat has a twining stem that grows 

as a vine whereas Japanese knotweed’s tall and erect stems do 

not need the support of other plants or objects (Chambers et al. 

1996) (Fig. 109). 

Figure 109: Comparing wild buckwheat (left) and Japanese knotweed 1 (right). 


japanese knotweed (Fallopia japonica) 

1..Plants.with.twining.stems...........................................................................Wild.buckwheat.(Fallopia convolvulus) 

1..Plants.with.tall,.erect.stems 

. 2..Plants.with.lobed.leaves 

. . 3..Leaves.deeply.and.palmately.lobed.(up.to.51cm.long)............Common.cow.parsnip.(Heracleum maximum) 

. . 3..Leaves.coarsely.and.unevenly.lobed.(up.to.152cm.long)........Giant.hogweed.(Heracleum mantegazzianum) 

. 2..Plants.with.entire.leaves 

. . 4..Lanced.to.elliptically-shaped.leaves...............................................Dock.(Rumex..spp.) 

. . 4..Triangularly-shaped.leaves................................................................Japanese.knotweed.(Fallopia japonica) 

BioloGy 


Japanese knotweed is native to Asia. It was introduced in both 

Europe and North America as an ornamental plant (Beerling 

et al. 1994). Japanese knotweed was introduced in Europe in 

the mid-19th century and in North America in 1877 (Forman &






........Subclass. Caryophyllidae 

..........Order. Caryophyllales 

............Family. Polygonaceae 

..............Genus. Fallopia. 

................Species. Fallopia japonica 


Kesseli, 2003). Canadian introductions 

were documented around the 1900’s 

and today Japanese knotweed is 

commonly found across Ontario 

(Bourchier & Van Hezewijk, 2010). 

hABitAt 

Japanese knotweed is often associated 

with nutrient-rich soils (Beerling et 

al. 1994) and is commonly found in 

both open and closed-canopy habitats 

such as old fields and forests. It has 

also been shown to be particularly 

invasive in wetlands and along stream 

and riverbanks (Maerz et al. 2005) 

(Fig. 110). 

Figure 110: Japanese knotweed in a riparian habitat 6 . 

reProduction 

Japanese knotweed is dioecious (i.e., with male and female 

individuals). Vegetative reproduction is important for this 

species, with the tiniest root fragments being capable of 

producing new plants (Beerling et al. 1994). Sexual reproduction 

and consequent seed production has been documented but the 

results are inconsistent. Some studies report the formation of 

empty seedpods or attribute high mortality rates to germinating 

seedlings (Locandro, 1973), while others report large numbers 

of highly viable seeds (Forman & Kesseli, 2003). More studies 

are needed to understand sexual reproduction as a dispersal 

mechanism for this exotic species. 

[ 103]

[ 104] 


liFe cycle 

Shoots begin to emerge and spread early in the spring and 

quickly grow in height. Flowering occurs from September 

through October with fruit development taking place soon 

thereafter (Beerling et al. 1994). New seedlings can flower in 

their first growing season (Forman & Kesseli, 2003). 


Japanese knotweed can form very dense patches. Over the 

years, dead stems and leaves build up a thick litter layer, which 

in combination with low light levels contribute to form an 

inhospitable environment for other plant species (Fig. 111). This 

effectively eliminates competition from native plant species and 

promotes Japanese knotweed invasion (Beerling et al. 1994). 

Figure 111: Japanese knotweed invasion with a visible accumulation of 

dead stems 1 . 

Japanese knotweed has a large and vigorous root system. The 

rhizomes form an extensive underground network that allows 

the plant to persist even if the above-ground portion is severely 

damaged. Japanese knotweed can rely on the reserves stored 

in the rhizome to boost its growth rate in the spring. This 

allows them to tower over native understory species to reach 

the sunlight (Beerling et al. 1994).



Japanese knotweed forms dense thickets that can effectively 

shade out native understory species (Fig. 112). The root systems 

grow outwards each year, sending up clonal shoots that can 

quickly form very dense stands. These stands displace native 

species and reduce available habitat for wildlife (Weber, 2003; 

Beerling et al. 1994). For instance, one study has shown that 

invasion by Japanese knotweed has a negative effect on frogs, 

which depend on terrestrial habitats such as forests and old 

fields for foraging and breeding. Indeed, areas with dense 

Japanese knotweed stands were found to be unsuitable breeding 

sites for green frogs (Maerz et al. 2005). 

Figure 112: A dense patch of Japanese knotweed 1 . 


Japanese knotweed disperses mainly vegetatively; even small 

pieces of root or stem can act as propagules and establish a new 

invasion. Yard wastes and soils containing plant fragments are 

thought to be the main vectors of Japanese knotweed’s dispersal 

into natural areas. As Japanese knotweed populations tend to 

grow in riparian areas, it has been assumed that waterways play 

a key role as a short and long distance transportation pathway 

(Beerling et al. 1994). Seed dispersal may also contribute to 

the spread of Japanese knotweed (Forman & Kesseli, 2003). 

However, more research is needed on the viability of Japanese 

knotweed seeds. 

[ 105]

[ 106] 




• Do not dump any type of yard waste in natural areas; 

• Ensure that soil being brought into a woodlot does not harbour any 

invasive seeds or plant fragments. Ask the distributor if the soil is 

classified as weed-free; 

• Do not accept or plant any unknown plant fragments in your garden. It 

may seem neighbourly to share gardening tips and plant slips. However, 

ensure that you know what you are planting before you do it. 


• Learn how to identify Japanese knotweed and ensure it is not growing 

in neighbouring areas; 

• Regularly monitor your woodlot paying attention to riparian areas where 

root fragments may have washed ashore. 


Excavation: Excavation is only feasible for very small 

populations. This should only be attempted when the population 

is recent and the root system is still relatively small. All root 

fragments should be removed because they can readily resprout. 

Excavation causes a great deal of soil disturbance. As 

such, it is recommended to plant desirable species in place of 

the Japanese knotweed to prevent invasion by other exotics 

(Stone, 2010). 

Cutting: Continuous cutting to eliminate the plant’s energy 

belowground can be an effective control strategy when dealing 

with large Japanese knotweed invasions. Every 2 to 3 weeks 

from spring to fall, cut the stems close to the ground, remove 

all the above-ground parts using a rake and bag everything. In 

areas with dense stands, re-sprouts may be controlled using 

a mower or weed whacker. Ongoing management for several 

years may be required to completely eliminate an invasion 

(Weston et al. 2005).


Herbicide application: Japanese knotweed invasions can be 

controlled using herbicides. A licenced exterminator will 

need to apply the herbicides, as effective systemic herbicides 

are needed to kill the robust root system. Several different 

application methods may be used, including foliar application 

and cut-stem methods. Foliar application may be more effective 

early in the season when the plants are still low to the ground. 

Japanese knotweed can reach 4m in height, which can make 

foliar herbicide applications a difficult task. As a result, cutting 

the stems may be a more practical method of control. Japanese 

knotweed stems can be quite dense, so cutting them will enable 

better accessibility. Herbicides can then be applied to the cross 

section of the stem (Remaley, 2005). 




Administer a foliar application in the spring when the plants 

are still short and the leaves are within reach. Monitor the area 

every couple of weeks and cut any re-sprouting individuals. 

Alternatively, a fall application can be done using the cut-stem 

method. The following spring, a foliar spray can be applied if resprouting 

is significant. In the event that only a few re-sprouts 

emerge, cutting or mowing can be employed every couple of 

weeks until the root reserves are exhausted (Remaley, 2005). 


Cut all stems close to the ground and remove the above-ground 

biomass. Cutting is recommended every 2 to 3 weeks from 

spring to fall. This will eventually eliminate any energy reserves 

stored in the roots. Management actions will be required for 

several years to completely eliminate an invasion (Weston et 

al. 2005). Digging up root crowns may speed up the control 

process. However, be prepared to plant native vegetation in 

areas where the soil has been disturbed. This will reduce the 

likelihood of other exotics establishing in the disturbed site. 

A plastic tarp may also be used on small areas where dense 

stands persist. Cut and remove all plant parts and cover the 

area with the plastic. The plastic will need to remain in place 

for several years to ensure that the roots die (Stone, 2010). 

[ 107]

[ 108] 


Extent.of. 

infestation 

Recommended. 

method.of. 

control 

Table 10: Management recommendations for Japanese knotweed (Fallopia japonica). 

Small.invasions.and.satellite. 

populations 


Excavation.and.cutting. Integrated.control: 

(Combination.of.physical.and.chemical. 

control). 

Timing Spring,.summer.&.fall. Cutting.and.hand-pulling.can.be.done. 

any.time.in.the.spring,.summer.and.fall.. 

Herbicide.application.should.occur.in.the. 

spring.or.fall.while.other.native.plants.are. 

dormant. 

Frequency.of. 

control 

Cutting.should.be.done.every.few. 

weeks.from.spring.to.fall. 

Length.of.control 2-3.years. 5+.years. 

Required. 

restoration 

Excavation.creates.soil.disturbance. 

that.benefits.exotic.species..Consider. 

planting.native.vegetation. 

Cut.stems.will.likely.re-sprout.and.require. 

frequent.clipping..Plan.to.control.every. 

couple.of.weeks. 

Restoration.is.not.feasible.until.the.invasion. 

is.under.control..Allow.native.vegetation.to. 

colonize.the.area.and.control.around.these. 

desirable.species.


5.1.6 

english ivy 



hardy english ivy 


109 

Figure 113: English ivy [ ] 

12. [ ]

[ 110] 



steM: English ivy has two growth forms. As an immature 

plant, the stem is a woody vine whereas mature plants look like 

a shrub (Reichard, 2000) (Fig. 114). 

Figure 114: Comparing the immature vining growth form 11 (left) to the mature shrub-like growth form 4 

(right) of English ivy. 

leAves: English ivy leaves are evergreen and grow in an 

alternate arrangement around the stem. The leaves on immature 

plants have 3 to 5 lobes whereas mature plants have lobeless 

cordate leaves (Fig. 115). They are dark green with lighter veins 

and a waxy coat (Kaufman & Kaufman, 2007). 

Figure 115: Comparing immature 12 (left) and mature 15 (right) English ivy leaves. 

Flowers: Clusters of yellowish-green flowers with 5 petals 

and 5 sepals are produced at the ends of the stems on mature 

plants (Swearingen et al. 2010) (Fig. 116).

5.1.6 ENGLISH IVY (Hedera helix) 

Fruit/seed: The fruit consists of a berry containing 3 to 5 

seeds. Berries grow in clusters and change from green to black 

when ripe (Miller, 2003; Gleason & Cronquist, 1963) (Fig. 117). 

heiGht: Vines can climb up to 28m high if they have a 

supporting structure (Miller, 2003) (Fig. 118). 

Figure 116: English ivy flowers 4 . Figure 117: Ripe English ivy 

berries 25 . 


Poison ivy (Toxicodendron radicans) 

Poison ivy usually grows as an erect or trailing shrub. However, 

it has been found growing as a climbing vine in parts of southern 

Ontario. As a vine, it may appear similar to English ivy with its 

leaves arranged in an alternate pattern. In contrast, poison ivy’s 

leaves are compound, containing three leaflets, whereas the 

leaves of English ivy are simple (Fig. 119). The fruits of poison 

ivy consist of white berries whereas the berries of English ivy 

are nearly black (Chambers et al. 1996). 

Figure 119: Comparing poison ivy 27 (left) to English ivy 15 (right). 

Figure 118: English ivy growing on 

a supporting tree 26 . 

[ 111 ]

[ 112] 


virginia creeper (Parthenocissus quinquefolia) 

Virginia creeper is a woody vine with similar dark coloured 

berries to those of English ivy. However, Virginia creeper has 

compound leaves, each with five leaflets (Fig. 120). The leaves 

of Virginia creeper are toothed whereas English ivy has leaves 

with smooth margins (Petrides, 1972). 

Figure 120: Comparing Virginia creeper 12 (left) to English ivy 12 (right). 

Grape (Vitis spp.) 

Frost grape (Vitis vulpina), summer grape (V. aestivalis), fox 

grape (V. lubrusca), and river bank grape (V. riparia) may be 

confused with English ivy. The easiest way to distinguish them 

from English ivy is to take a close look at the leaf margins. 

English ivy has smooth leaf margins whereas all of these grape 

species have toothed margins (Fig. 121). See the key to help 

differentiate between grape species (Petrides, 1972). 

Figure 121: Comparing the leaves of a species of grape 1 (left) to those of English ivy 1 (right).


canada moonseed (Menispermum canadense) 

Canada moonseed may be confused with English ivy. Both 

species are trailing, woody vines with alternately arranged 

simple leaves with smooth edges (Fig. 122). One clear difference 

between these two species is that the petioles of Canada 

moonseed are attached to the base of the leaf whereas those 

of English ivy are attached to the end (Petrides, 1972) (Fig. 123). 

Figure 122: Comparing the leaves of Canada moonseed 1 (left) to those 

of English ivy 12 (right). 

Figure 123: Canada moonseed 

petiole attachment 20 . 

A key to plants with climbing stems that may be 

confused with english ivy (Hedera helix) 

1..Plants.with.compound.leaves 

. 2..Leaves.with.three.leaflets....................................................................... Poison.ivy.(Toxicodendron radicans) 

. 2..Leaves.with.five.leaflets.......................................................................... Virginia.creeper.(Parthenocissus quinquefolia) 

1..Plants.with.simple.leaves 

. . 3..Leaves.with.toothed.(serrate).edges 

. . . 4..Leaves.without.lobes...................................................................... Frost.grape.(Vitis vulpina) 

. . . 4..Leaves.with.lobes 

. . . . 5..Leaf.undersides.are.white........................................................ Summer.grape.(Vitis aestivalis) 

. . . . 5..Leaf.undersides.not.white 

. . . . . 6..Leaf.undersides.densely.hairy........................................... Fox.grape.(Vitis lubrusca) 

. . . . . 6..Leaf.undersides.slightly.hairy.along.veins..................... River.bank.grape.(Vitis riparia) 

. . 3..Leaves.with.smooth.(entire).edges 

. . . . . . 7..Petioles.attached.to.leaf.undersides........................... Canada.moonseed.(Menispermum canadense) 

. . . . . . ....(peltate.leaves) 

. . . . . . 7..Petioles.attached.to.leaf.margins................................. English.ivy.(Hedera helix) 

[ 113]

[ 114 ] 







........Subclass. Rosidae 

..........Order. Apiales 

..........Family. Araliaceae 

.............Genus. Hedera 

...............Species. Hedera helix 

BioloGy 


English ivy has been used as an 

ornamental ground cover since the 

early 1700’s when it was introduced 

from Europe, western Asia and 

northern Africa (Swearingen et al. 

2010). Today it is still commonly sold 

as a garden plant throughout Ontario 

(Fig. 124). 

hABitAt 

English ivy can grow in a wide range 

of habitats in either full shade or 

sunlight (Swearingen et al. 2010). 

However, it thrives in open-canopy 

Figure 124: English ivy being sold as a ground cover species 1 . 

forests (Miller, 2003). Deciduous forests are a prime habitat 

for English ivy invasion as their evergreen leaves benefit from 

full access to the sun before the tree canopy establishes in the 

spring (Fig. 125). Populations are often found in forests close to 

urban areas as they frequently escape from gardens (Reichard, 

2000).


Figure 125: English ivy growing in an open-canopy forest 25 . 

reProduction 

English ivy spreads vegetatively and by seeds. The species 

has a juvenile and a mature life stage. Juvenile plants grow as 

woody vines and do not produce flowers or fruits (Reichard, 

2000). Maturity can take up to ten years and is often initiated 

by access to full sunlight conditions. In the mature stage the 

plant takes on a shrub-like growth form and begins to produce 

flowers and seeds. Stem fragments are able to root and produce 

new plants (Swearingen & Diedrich, 2006). 

liFe cycle 

English ivy is a perennial evergreen vine. Mature plants flower 

throughout the summer and produce fruit in the fall. The pale 

green berries turn dark purple as they ripen and may stay on 

the plant throughout the winter (Miller, 2003). 

[ 115 ]

[ 116] 



The vining habit of English ivy coupled with its fast vegetative 

spread gives this species a competitive edge over other native 

plants. It can quickly climb over any supporting structure 

towards sunlight, smothering other plants and limiting their 

access to sunlight (Dlugosch, 2005). 

The evergreen life-strategy allows English ivy to photosynthesize 

and grow earlier than native deciduous plants (Fig. 126). This 

jumpstart growth in the spring gives English ivy a competitive 

edge over other plants. As the forest canopy closes English ivy 

has the added benefit of shade tolerance. Therefore, not only 

can this plant thrive in shaded environments but it can also 

quickly climb to reach more preferable light levels (Swearingen 

& Diedrich, 2006). 

Figure 126: The evergreen leaves of English ivy 15 . 


English ivy can form a dense carpet on the forest floor. They 

climb over native understory species, depriving them of sunlight 

and, as a result, exclude native species (Fig. 127). Germinating 

seedlings cannot compete with such a dense vegetative ground 

cover, which may cause a decrease in hardwood recruitment. 

With little to no recruitment, the species composition of the 

forest may shift (Dlugosch, 2005). 

English ivy can climb over supporting structures such as trees 

and reach heights of 28m (Fig. 128). As these vines climb up 

the trunks of hardwood trees and onto their branches they


effectively prevent the sunlight from reaching the tree’s leaves, 

causing a decrease in its vigor (Miller, 2003). In heavy invasions, 

English ivy has the ability to kill the host tree (Swearingen & 

Diedrich, 2006). These vines also create added weight that 

can cause trees to collapse during high winds or snow storms 

(Swearingen et al. 2010). The vining habit of English ivy creates 

pockets of water close to the tree trunks, which often causes 

fungal growth and decay (Kaufman & Kaufman, 2007). 

Figure 127: A carpet of English ivy in the forest 

understory 15 . 

English ivy can carry a generalist bacterial plant pathogen 

(Xylella fastidiosa), which is the causing agent of leaf scorch. This 

pathogen affects the xylem of hardwood species. Infected trees 

show signs of scorched leaves and stunted growth (McElrone et 

al. 1999) (Fig. 129). 

Figure 129: Symptoms of bacterial leaf scorch 90 . 

Figure 128: English ivy climbing into the forest 

canopy 16 . 

[ 117 ]

[ 118] 



English ivy can be purchased throughout Ontario and is 

commonly found as an ornamental ground cover in urban 

gardens (Miller, 2003). The dark coloured berries on mature 

English ivy plants are attractive to birds. These fruits contain 

glycosides and other compounds which cause the seeds to 

quickly pass through the digestive track or cause the birds 

to regurgitate. In this way seeds can be dispersed over long 

distances and into natural areas. Some spread may also be due 

to the dumping of yard wastes into natural areas as new plants 

can easily grow from stem and root fragments (Swearingen et 

al. 2010). 



• Do not dump yard wastes in or close to forested areas; 

• English ivy is considered an invasive plant and should not be 

planted in the garden. Use native alternatives or species that are not 

considered invasive (Fig. 130). 


• Learn how to identify English ivy; 

• Regularly monitor the woodlot for invasive species; 

• Talk with neighbours about what they grow in their gardens and the 

threats associated with growing invasive plants; 

• Ensure that yard wastes are properly disposed of. 

Figure 130: English ivy is a common ground cover and border plant for gardens 25 .



Hand-pulling: Small patches and satellite populations can 

be controlled by hand-pulling. From a kneeling position, 

individual vines should be uprooted as far as one can reach. 

Continue moving and uprooting vines until they all have been 

removed from the area (Soll, 2005). Be careful to collect as much 

of the root system as possible because even the smallest root 

fragments have the ability to re-sprout. Vines should be bagged 

and disposed of in an appropriate landfill. Monitor the area for 

approximately 2 to 3 years to ensure that no re-sprouting has 

occurred (Swearingen & Diedrich, 2006). 

Herbicide application: Apply herbicide with a surfactant as 

a foliar spray to English ivy running along the ground. The 

surfactant is needed to help the herbicide penetrate the waxy 

layer of English ivy leaves. Apply herbicide to leaves until just 

wet and try to avoid any dripping (Miller, 2003). For English ivy 

that is climbing up trees, it is best to cut the vine and apply the 

herbicide to the stump (Swearingen & Diedrich, 2006). If done 

properly, herbicide treatments can eliminate the majority of 

English ivy invasion in the first treatment. Follow-up treatments 

will be needed to completely eradicate the population (Soll, 

2005). 



options #1: with chemical control 

Plan to apply herbicide in the fall when other native vegetation 

is dormant. Choose a period when rain is not forecasted for at 

least several days. Perform spot herbicide application in dense 

patches of English ivy while hand-pulling scattered individuals 

or patches close to desirable vegetation. Hand-pulling can be 

done before or after herbicide application depending on the 

severity of the invasion. In areas with dense patches of English 

ivy and only a small number of native plants, herbicides can be 

applied in the fall, being careful to spray around any desirable 

vegetation. English ivy growing beside desirable vegetation 

should not be sprayed and should be hand-pulled at a later 

date. Hand-pulling any remaining English ivy plants can be 

scheduled for the following spring (Soll, 2005). 

Depending on the effectiveness of the initial control measure, 

follow-up treatments may include a second herbicide application. 

It may take several months before any effects from the initial 

herbicide application are visible. If dense patches remain, a 

follow-up foliar spray or spot application may be needed (Soll, 

2005). For small patches, manual removal of the entire plant 

is appropriate. Carefully pull up all roots and bag the plant 

[ 119 ]

[ 120] 


Table 11: Management recommendations for English ivy (Hedera helix). 


Recommended. 


Manual.control:.hand-pulling. Integrated.control:.hand-pulling.&. 

herbicide.application. 

Timing Spring,.summer.and.fall. Spring.and.fall. 

Disposal Bag.all.parts.of.the.plant.and.dispose.at. 

an.appropriate.landfill. 

Frequency.of. 

control 

Initial.pull.should.remove.as.many.plants. 

and.plant.fragments.as.possible..Return. 

several.times.per.year.to.pull.any.missed. 

plants.or.re-sprouts. 


Required. 

restoration 

for disposal to ensure that roots do not re-sprout. Monitor the 

area for approximately 3 to 5 years after the initial treatment 

(Swearingen & Diedrich, 2006). Planting native vegetation in 

the infested area after the initial treatment is recommended. 

This will promote natural re-growth and may increase natural 

resistance against the establishment of invasive plants (Soll, 

2005). 


Hand-pulling should be scheduled for the spring with a follow-up 

in the fall. However, most English ivy plants should be removed 

in the first hand-pulling. Be sure to collect as much of the root 

system as possible. Do not leave any root fragments behind 

as they can easily re-sprout. Being thorough during the first 

hand-pulling will make future management activities easier or 

potentially unnecessary (Swearingen & Diedrich, 2006). Vines 

should be bagged and disposed of in an appropriate landfill. 

Return in the fall to pull any remaining English ivy plants and 

monitor the area for approximately 3 to 5 years after the initial 

control (Miller, 2003). 

Plant.native.species.in.areas.where.handpulling.creates.soil.disturbance. 

Bag.all.hand-pulled.plants.and.dispose. 

at.an.appropriate.landfill. 

Manual.removal.in.the.spring.and. 

herbicide.application.in.the.fall.may.be. 

required.for.up.to.3.years.depending.on. 

extent.of.the.infestation..As.infestation. 

gets.smaller.manual.removal.should. 

replace.chemical.control. 

Plant.native.species.in.areas.where.handpulling.creates.soil.disturbance.


5.1.7 

himalayan Balsam 

(Impatiens glandulifera) 


Purple jewelweed, Policeman’s helmet, indian balsam, ornamental 

jewelweed 


121 

Figure 131: Himalayan balsam [ ] 

1 [ ]

[ 122] 



leAves: The leaves of Himalayan balsam are often found 

whorled around the stem in groups of three (Fig. 132). Some 

leaves may also be found as pairs in an opposite arrangement. 

The leaf edges are sharply toothed and lanceolate or elliptical 

in shape (Beerling & Perrins, 1993). 

Figure 132: Himalayan balsam leaves 1 . 

Flowers: The flowers of Himalayan balsam are irregular 

and trumpet-shaped (Fig. 133). They range in colour from dark 

pink to white and grow in clusters of 3 to 12 flowers (Beerling 

& Perrins, 1993) (Fig. 134). 

Figure 133: Himalayan balsam flowers 1 Figure 134: Colour variation in the flowers of Himalayan balsam 6 . 

Fruit/seed: Seeds are found within green capsules (Fig. 135). 

When ripe, these capsules burst open at the slightest touch to 

disperse 4 to 16 seeds (Beerling & Perrins, 1993). 

steM: The stems are reddish-green and hollow (Fig. 136). They 

can grow as large as 5cm in diameter (Beerling & Perrins, 1993). 

heiGht: In North America, this herbaceous herb can grow 

up to 3m tall, towering over other native vegetation in the 

understory (Tabak & von Wettberg, 2008) (Fig. 137).

5.1.7 HIMALAYAN BALSAM (Impatiens glandulifera) 

Figure 135: Himalayan balsam seedpods 1 . Figure 136: The hollow stem of Himalayan balsam 1 . 

Figure 137: Himalayan balsam towering over native vegetation 1 . 

[ 123]

[ 124] 



Himalayan balsam can be confused with native members of 

the “touch-me-not” family that have irregular flowers and 

“exploding” seedpods. The easiest way to identify Himalayan 

balsam is to look at the arrangement of the leaves on the stem. 

Himalayan balsam is the only species of Impatiens in Ontario 

with whorled or opposite leaves. 

spotted touch-me-not (Impatiens capensis) 

Spotted touch-me-not is a native plant in Ontario commonly 

found in wet areas. The flowers have a similar shape to those 

of Himalayan balsam. However, they are orange with tiny red 

spots (Fig. 138). The leaves have irregular toothed margins and 

an alternate arrangement along the stem (Dickinson et al. 2004) 

(Fig. 139). 

Figure 138: Comparing the flowers of 

spotted touch-me-not 1 (top) to those 

of Himalayan balsam 1 (bottom). 

Pale touch-me-not (Impatiens pallida) 

Figure 139: Comparing the leaves of 

spotted touch-me-not 1 (top) to those 

of Himalayan balsam 1 (bottom). 

Pale touch-me-not is a native species with similar irregular 

flowers to those of Himalayan balsam. These flowers are pale 

yellow in contrast to the pink or white flowers of Himalayan 

balsam (Fig. 140). The leaves have an alternate arrangement 

along the stem with regular toothed margins (Dickinson et al. 

2004).


Figure 140: Comparing the flowers of pale touch-me-not 15 (left) to those of Himalayan balsam 1 (right). 


himalayan balsam (Impatiens glandulifera) 

1..Plants.with.irregular.flowers 

. 2..Plants.with.orange.or.yellow.flowers 

. . 3..Orange.flowers.with.red.spots............................................................. Spotted.touch-me-not.(Impatiens capensis) 

. . 3..Pale.yellow.flowers.................................................................................. Pale.touch-me-not.(Impatiens pallida) 

. 2..Plants.with.pink,.purple.or.white.flowers............................................. Himalayan.balsam.(Impatiens glandulifera) 

1..Plants.without.flowers.present 

. . . . 4..Alternate.leaves 

. . . . . 5...Irregular.toothed.margins...................................................... Spotted.touch-me-not.(Impatiens capensis) 

. . . . . 5..Regular.toothed.margins........................................................ Pale.touch-me-not.(Impatiens pallida) 

. . . . 4..Whorled.or.opposite.leaves......................................................... Himalayan.balsam.(Impatiens glandulifera) 







..........Order. Geraniales 

............Family. Balsaminaceae 

..............Genus. Impatiens 

................Species. Impatiens glandulifera 

BioloGy 


Himalayan balsam is native to the 

western Himalayas. It has become 

an invasive plant in North America, 

Europe and New Zealand where it 

was most likely introduced as an 

ornamental garden plant due to its 

large, attractive flowers. The first 

records of Himalayan balsam in 

North America are from Connecticut 

in 1883 (Tabak & von Wettberg, 

2008). It has since been introduced to 

Ontario as a garden plant. 

[ 125]

[ 126] 


hABitAt 

Himalayan balsam can grow in a wide variety of habitats. It 

has become a particularly troublesome invader along riparian 

areas, wetlands and forest understories in Europe (Tabak & von 

Wettberg, 2008), where dense stands have been reported in both 

mixed and deciduous forests. In Ontario, Himalayan balsam 

is prevalent in riparian areas (Fig. 141) and has the potential 

to become a problem 

in hardwood forests 

where localized disturbances 

allow this 

species to establish 

and spread (Perrins et 

al. 1993; Ammer et al. 

2011). 

reProduction 

Himalayan balsam reproduces 

by seed. The 

flowers are able to selfpollinate. 

However, this 

is rarely needed as the 

large colourful flowers 

are attractive to insect Figure 141: Himalayan balsam growing alongside a stream 

pollinators such as bees 

and wasps (Bartomeus et al. 2010). Individual plants have the 

potential to produce an average of 800 seeds. These seeds are 

contained in pods that “explode” upon maturity when touched, 

allowing seeds to disperse several metres away from the parent 

plant. Seeds can survive in the soil for up to 18 months. No 

vegetative reproduction has been documented for Himalayan 

balsam (Perrins et al. 1993; Beerling & Perrins, 1993). 

29 . 

liFe cycle 

Himalayan balsam is an annual plant (i.e., it completes its life 

cycle in one growing season). Seedlings germinate and emerge 

in the early spring and quickly grow into tall flowering plants. 

Flowering begins in July and the formation of seedpods can 

occur as early as mid-July (Beerling & Perrins, 1993). Seedpods 

ripen and become sensitive to the touch from August to October 

(Ammer et al. 2011). 


Himalayan balsam`s high reproductive potential contributes 

to its success as an invader. Prolific seed production allows 

populations to quickly increase in size while self-pollination 

allows for increased dispersal and establishment of this 

species. Transportation of a single seed to a new area can result


in an invasion because Himalayan balsam does not solely rely 

on cross-pollination (Bartomeus et al. 2010; Perrins et al. 1993). 

Factors that give Himalayan balsam a competitive advantage 

are synchronous seed germination and high vegetative growth 

rate. Seeds germinate in the early spring and all within a few 

days of each other. After seed germination the seedlings gain 

height and biomass in a very short amount of time. Native 

plants in the forest understory generally have a slower growth 

rate and do not exhibit synchronous germination. The presence 

of a large number of tall plants prevents native seedlings from 

acquiring the necessary amounts of light and energy to grow 

(Fig. 142). Very few native seedlings can survive under such low 

light conditions in the early spring (Beerling & Perrins, 1993; 

Andrews et al. 2009). 


European habitats have been experiencing the negative effects 

of Himalayan balsam invasion since the 1850’s, while its first 

introduction into North America occurred approximately 30 

years later (Tabak & von Wettberg, 2008). Thus, most studies 

documenting the ecological impacts of Himalayan balsam 

have been done in Europe (Perrins et al. 1993; Maule et al. 

2000; Andrews et al. 2009). These studies can help us predict 

the effects Himalayan balsam may have on hardwood forests 

in Ontario. As with other invasive plants, Himalayan balsam 

forms dense stands (Fig. 143) that compete and exclude native 

vegetation from the area (Andrews et al. 2009). This competition 

may reduce forest biodiversity and have an impact on hardwood 

regeneration (Pyšek & Prach, 1995). However, more research is 

needed to fully understand the impacts of Himalayan balsam 

on North American hardwood stands. 

Figure 142: Himalayan balsam attaining great height 4 . Figure 143: A dense stand of Himalayan balsam 1 . 

[ 127]

[ 128] 



Dispersal is initiated when the seedpods burst open, expelling 

seeds within several metres of the parent plant. Long-distance 

dispersal can occur when plants are close to streams or rivers. 

The seeds are buoyant and can be carried downstream (Tabak 

& von Wettberg, 2008). Humans are probably the main source 

of long-distance seed dispersal. Seeds can easily get stuck in 

clothing folds, shoes or on vehicle tires. Some dispersal may 

also be caused by the use of Himalayan balsam as a garden 

plant (Perrins et al. 1993). 



• Himalayan balsam can easily become established in areas where soil 

has been disturbed. Limit soil disturbing activities in the woodlot 

and plant native vegetation in areas where soil disturbance is 

unavoidable; 

• If a river or stream runs through the woodlot, ensure that populations 

of Himalayan balsam are controlled upstream to prevent seed 

dispersal; 

• Talk with neighbours about the potential consequences of using 

Himalayan balsam as a garden plant and suggest some native 

alternatives. 


• Learn how to identify Himalayan balsam; 

• Monitor the woodlot frequently paying attention to disturbed areas 

and along waterways; 

• Become familiar with Himalayan balsam in neighbouring gardens. 


Hand-pulling: Although hand-pulling may prove to be time 

consuming, the shallow root system of these plants makes 

pulling relatively easy. Hand-pulling is appropriate for areas 

where Himalayan balsam is mixed with desirable vegetation. 

Take hold of the stem as close to the ground as possible to 

prevent the stem from breaking. The root systems of some of 

the larger plants may prove difficult to remove. The roots may 

be dug up using a hand trowel to prevent any re-sprouting (Kelly 

et al. 2008). Himalayan balsam has a very shallow root system


which allows plants to be easily pulled from the soil. Even with 

a shallow root system, hand-pulling Himalayan balsam will 

create soil disturbance. Consider planting native vegetation 

to prevent re-colonization or invasion by other problematic 

species (Kelly et al. 2008). 

Cutting and mowing: Cutting or mowing in areas with dense 

populations of Himalayan balsam can prevent these plants 

from producing seed. Using a weed whacker on dense patches 

of Himalayan balsam is a relatively quick method of removal. 

Make sure the plants are cut as close to the ground as possible, 

ideally below the lowest node, to prevent re-sprouting. This 

can be done several times between April and June before 

the plants have produced seedpods as this will scatter seeds 

thereby increasing the area of invasion (Havinga, 2000). Be sure 

to schedule a cutting towards the end of June to prevent late 

growing plants from reaching the fruiting stage. Ensure that 

stems are cut as close to the ground as possible to prevent resprouting 

(Kelly et al. 2008). 

Herbicide application: Chemical control by a licenced 

exterminator is another way of managing large invasions of 

Himalayan balsam. Herbicides should be applied as a foliar 

spray after flowering but before seedpod development. Any 

plants that are sprayed after flowering should not have enough 

energy to produce viable seed. This approach should effectively 

prevent seed production and result in a smaller population the 

following year (Kelly et al. 2008). 




Control efforts should be done between April and June before 

seedpods begin to develop. This timing is important because 

seedpods will explode at the slightest touch, releasing seeds 

and contributing to the seed bank (Kelly et al. 2008). Seeds are 

only known to survive in the soil for a maximum of 18 months 

so control measures should only be needed for 2 consecutive 

years (Beerling & Perrins, 1993). 

In areas with large populations of Himalayan balsam a 

combination of manual and chemical control may be required. 

Herbicides can be applied as foliar sprays to dense patches. 

In areas where herbicides cannot be used, such as areas close 

to water, cutting down large invasions can help reduce seed 

production. Hand-pulling should be employed in areas where 

Himalayan balsam is intermingled with desirable vegetation. 

Consider planting native species in areas where hand-pulling 

has created soil disturbance or where control has left bare 

areas (Kelly et al. 2008; Havinga, 2000). 

[ 129]

[ 130] 



The best way to control small invasions of Himalayan balsam 

is to hand-pull individual plants. Hand-pulling should be done 

between April and June before seedpods begin to develop. 

This timing is important because seedpods will explode at the 

slightest touch, releasing seeds and contributing to the seed 

bank (Kelly et al. 2008). If timed appropriately, only one pull 

will need to be scheduled per year. By waiting until plants 

have begun to produce flowers, any seedlings that germinate 

thereafter should not have enough time to mature and produce 

seed before winter (Ammer et al. 2011). Pulled plants should 

be bagged and disposed of in an appropriate landfill or left 

to decompose on a tarp. In areas with large populations, use 

a weed whacker or lawn mower. Cuttings should be done 

several times per season to prevent any addition to the seed 

bank. After several years, the seed bank should be exhausted 

and populations should start to diminish (Kelly et al. 2008; 

Havinga, 2000). 

Table 12: Management recommendations for Himalayan balsam (Impatiens glandulifera). 


populations 

Recommended. 



Manual.control:.hand-pulling. Manual.control,.cutting/mowing,.handpulling,.chemical.control,.and.herbicide. 

application. 

Timing April-June. Manual.control:.April-June. 

Chemical.control:.May-early.June. 

(immediately.after.flowering). 

Disposal Bag.all.parts.of.the.plant.and.dispose. 

at.an.appropriate.landfill.or.leave.to. 

decompose.on.a.tarp. 

Frequency.of. 

control 

Initial.pull.should.remove.as. 

many.plants.as.possible..If.timed. 

appropriately,.only.one.pull.per.year. 

will.be.needed. 


Required. 

restoration 

Plant.native.species.in.areas.where. 

hand-pulling.creates.soil.disturbance. 

Plant.material.may.be.left.on.site. 

Cutting.or.mowing.should.be.down.several. 

times.per.year.between.April.and.June. 

Herbicides.may.be.applied.once.per.year. 

between.May.and.early.June. 

Plant.native.species.in.areas.with.soil. 

disturbance.or.where.controlled.patches. 

have.left.bare.areas


5.1.8 

common Buckthorn 

(Rhamnus cathartica) 


european buckthorn, carolina buckthorn, european waythorn, 

hart’s thorn 


131 

Figure 144: Common buckthorn [ ] 

1 [ ]

[ 132] 



leAves: Common buckthorn leaves are usually found in an 

opposite arrangement but at times can be alternate. They are 

oval in shape with toothed margins (Fig. 145). Each leaf has 3 to 

4 pairs of veins that curve towards the leaf tip (Wieseler, 2005; 

Kershaw, 2001). 

Figure 145: Common buckthorn leaves 1 . 

Flowers: Common buckthorn is dioecious, which means 

that each plant produces either male or female flowers (Fig. 

146). Flowers are greenish-yellow in colour with 4 petals. They 

are arranged in a cluster at the leaf axils (Kershaw, 2001). Each 

cluster will often have 2 to 6 flowers (Wieseler, 2005). 

Figure 146: Pistillate or female flowers1 (left) and staminate or male 

flowers1 (right) of common buckthorn. 

Fruit/seed: The fruit is a berry containing approximately 

four seeds. As the berries mature they change from green to 

red to black (Fig. 147). They are arranged in clusters (Kershaw, 

2001). 

trunk & BrAnches: The bark has characteristic spots 

called lenticels (Fig. 148). Branches and twigs are commonly 

spine-tipped (Kershaw, 2001).

Figure 147: Immature green (top) and ripe black 

(bottom) common buckthorn berries 1 . 

5.1.8 COMMON BUCKTHORN (Rhamnus cathartica) 

Figure 148: Common buckthorn trunk with characteristic 

lenticels 1 (top) and a spine-tipped twig 1 (bottom). 

heiGht: Common buckthorn is a multiple-stemmed shrub 

or single-stemmed small tree that can reach up to 6m tall 

(Kershaw, 2001) (Fig. 149). 

Figure 149: Common buckthorn growing as a small tree 1 . 

[ 133]

[ 134] 



Glossy buckthorn (Frangula alnus) 

Glossy buckthorn has similar leaf venation patterns to that of 

common buckthorn. However, the leaves have smooth edges 

and 5 to 10 pairs of veins. This is in contrast with common 

buckthorn, whose leaves have toothed edges and 2 to 4 veins 

per side (Fig. 150). Other differences include the glossy leaves 

and spineless twig tips of glossy buckthorn (Kershaw, 2001). 

Figure 150: Comparing the leaves of glossy buckthorn 14 (left) to those of common buckthorn 1 (right). 

red-osier dogwood (Cornus sericea) 

The leaves of red-osier dogwood have a similar venation pattern 

to those of common buckthorn. However, the leaf edges are 

smooth and not toothed as those of common buckthorn (Fig. 

151). Red-osier dogwood has a characteristic bright red stem 

that is distinctive from other shrubs (Kershaw, 2001). 

Figure 151: Comparing the leaves of red-osier dogwood 1 (left) to those of common buckthorn 1 (right). 

Alternate-leaved dogwood (Cornus alternifolia) 

As with red-osier dogwood, the leaf edges of alternate-leaved 

dogwood are smooth, which is in contrast to the toothed edges 

of common buckthorn’s leaves. Common buckthorn has 3 to 

4 vein pairs while alternate-leave dogwood has 5 to 6 vein 

pairs (Fig. 152). Although the leaves of common buckthorn are 

usually found in an opposite arrangement, they can at times 

have the same alternate arrangement typical of alternate-leaved 

dogwood (Farrar, 1995)


Figure 152: Comparing the leaves of alternate-leaved dogwood 1 (left) to those of common buckthorn 1 (right). 

Alder-leaved buckthorn (Rhamnus alnifolia) 

A close relative of common buckthorn, alder-leaved buckthorn, 

is very similar in appearance. Both species have leaves with 

toothed edges and arcuate venation. However, common 

buckthorn has 3 to 4 vein pairs whereas alder-leaved buckthorn 

has 6 to 7 vein pairs (Fig. 153). The best way to distinguish these 

species from one another is to count their vein pairs (Chambers 

et al. 1996). 

Figure 153: Comparing the leaves of alder-leaved buckthorn 14 (left) to those of common buckthorn 1 (right). 

A key to woody plants with leaves similar to 

common buckthorn (Rhamnus cathartica) 

1..Leaves.with.toothed.edges 

. 2..Leaves.are.glossy....................................................................................Glossy.buckthorn.(Frangula alnus) 

. 2..Leaves.are.not.glossy 

. . 3..Leaves.in.an.opposite.arrangement............................................Red-osier.dogwood.(Cornus sericea) 

. . 3..Leaves.in.an.alternate.arrangement...........................................Alternate-leaved.dogwood.(Cornus alternifolia) 

1..Leaves.with.smooth.edges 

. . . 4..Leaves.with.6-7.pairs.of.veins...................................................Alder-leaved.buckthorn.(Rhamnus alnifolia) 

. . . 4..Leaves.with.3-4.pairs.of.veins...................................................Common.buckthorn.(Rhamnus cathartica) 

[ 135]

[ 136] 








..........Order. Rhamnales 

............Family. Rhamnaceae 

..............Genus. Rhamnus 

................Species. Rhamnus cathartica 

BioloGy 


Common buckthorn is native to 

Europe and Asia. It was introduced 

into North America in the early 1800’s 

as an ornamental shrub (Knight et al. 

2007) and it was highly valued for its 

aesthetic appearance as an hedge plant 

(Kershaw, 2001). Common buckthorn 

escaped from cultivation and spread 

across southern Ontario through 

the 1900’s (Kurylo et al. 2007). After 

being discovered as an alternate host 

for various crop pests such as oat 

crown rust (Puccinia coronata; Fig. 

154), a ban was put on its importation 

and sale within Canada (CFIA, 2008). 

Figure 154: Crown rust on an oat leaf 30 . 

hABitAt 

Common buckthorn can be found occupying forest edges and 

open areas in its native range (Knight et al. 2007). In North 

America it can be found in both open and closed habitats such 

as fields, forests and disturbed sites (Mascaro & Schnitzer, 

2007). Common buckthorn is a shade tolerant species that can 

readily invade forest understories (Kurylo et al. 2007; Knight et 

al. 2007) (Fig. 155).


Figure 155: Common buckthorn invading the edge of a forest 1 . 

reProduction 

Common buckthorn is dioecious, which means that each plant 

produces either male or females flowers. Only female plants 

produce fruit and seed (Wieseler, 2005). It may take 9 to 20 

years before common buckthorn can reproduce. After that it 

can reproduce every year. Reproduction is sexual and flowers 

require insect pollinators (Knight et al. 2007). Seeds can persist 

for at least 2 years in the soil after common buckthorn has been 

removed from an area (Delanoy & Archibold, 2007). 

liFe cycle 

Seeds usually disperse in late fall and remain dormant until 

germination the following summer. The leaves of common 

buckthorn are deciduous but they generally appear earlier and 

fall later than those of native trees and shrubs. Leaves generally 

emerge in April and May followed by flowering in May and June. 

Fruits ripen in September and may remain on the plant during 

the winter months (Knight, 2005) (Fig. 156). 

Figure 156: Common buckthorn berries in January 14 . 

[ 137]

[ 138] 



A fast growth rate has been shown to be advantageous for 

common buckthorn in North America. In the early spring it 

produces leaves much earlier than co-occurring native plants. 

This enables exposure to full sunlight before the forest 

canopy closes, resulting in a jump-start in energy reserves. 

This, in conjunction with keeping its leaves longer than their 

native competitors in the fall, results in an overall longer 

photosynthetic period, which may contribute to its success in 

the forest understory (Knight et al. 2007). 

Emodin is a secondary compound found in the leaf tissues and 

unripe berries of common buckthorn. Emodin is thought to 

play a role in protection against pathogen attack and herbivory. 

Emodin present in unripe berries may make them unpalatable to 

birds and other wildlife early in the season. As fruits ripen, their 

concentration of emodin decreases. This allows fruits to mature 

before they are consumed by wildlife which is thought to play 

an important role in seed viability and dispersal (Izhaki, 2002). 


Common buckthorn is a competitive shrub that is capable 

of becoming a dominant understory species (Knight et al. 

2007) (Fig. 157). Early leaf production and a fast growth rate 

allow these shrubs to tower over and smother understory 

plants. Dense shade from established thickets creates inade- 

quate growing conditions 

for native species, thereby 

reducing hardwood recruitment 

(Delanoy & Archibold, 

2007; Mascaro & Schnitzer, 

2007). 

Common buckthorn invasions 

can change soil characteristics 

and forest floor communities. 

The leaves contain high 

levels of nitrogen which is 

incorporated in the forest floor 

as they decompose. Studies 

have shown that earthworms 

are attracted to sites invaded 

by common buckthorn. 

Figure 157: A thicket of common buckthorn 

saplings in a forest understory 

High levels of earthworm 

colonization can lead to increased litter decomposition rates and 

thus to major changes in the dynamics of both food webs and 

nutrient cycling (Heneghan et al. 2007; Knight et al. 2007). 

1 .


As common buckthorn becomes established in the forest 

understory it can effectively exclude co-occurring native 

shrubs, resulting in large monospecific thickets (Knight et 

al. 2007). Songbirds may be forced to nest in these thickets 

due to the displacement of native shrubs. A study by Schmidt 

and Whelan (1999) documented increased predation rates on 

American robin (Turdus migratorius) nests found in common 

buckthorn thickets as compared to those nests found on native 

shrubs. More studies are needed to determine the impact of 

common buckthorn on songbird populations. 


Common buckthorn spreads via the dispersal of seeds. These 

seeds are borne in fruits that either drop to the ground or are 

eaten by wildlife. The berries can float and may be washed 

away by flood waters or carried down streams and rivers. Since 

birds can fly long distances before seeds pass through their 

digestive tracks, they are presumably the primary means of 

common buckthorn dispersal into hardwood forests (Gale, 

2000). 

The importation or sale of common buckthorn is prohibited in 

Canada due to its ability to act as an alternate host for oat crown 

rust (Puccinia coronata) and other crop pests such as potato 

and soybean aphids (CFIA, 2008; Knight, 2005). Although it can 

no longer be sold as an ornamental shrub, humans may still act 

as dispersal agents when seeds get stuck on shoes and clothing 

or in the treads of tires. 




washing all shoes and tires before entry; 

• Increase awareness in your community about the consequences of 

common buckthorn invasions. Promote management to help control 

common buckthorn. 


• Learn how to identify common buckthorn; 

• Regularly monitor the woodlot for new species and pay particular 

attention to roads and trails where invasive plants can easily establish. 

[ 139]

[ 140 ] 



Hand-pulling: Common buckthorn can appear as a shrub with 

multiple stems or as a tree that can reach 6m in height (Kershaw, 

2001). Hand-pulling is only practical for small saplings. Wet soil 

makes hand-pulling easier. As such, it is best to plan for manual 

control after a heavy rainfall. Small seedlings that are less than 

1m tall can be pulled but gloves should be worn to prevent 

injury from the thorny branches (Gale, 2000). For saplings that 

are larger than 1m in height, mechanical levers such as root 

wrenches will be needed to pry up the root system from the 

soil. This may prove to be very labour intensive and should 

only be attempted for small invasions. 

Cutting and excavation: Large trees may have to be cut and 

the roots dug out using a shovel (Gale, 2000). The root systems 

must be removed because shoots will emerge from root crowns 

and stumps (Pergams & Norton, 2006). Since mechanical 

removal will cause significant soil disturbance, restoration 

efforts will be required. Fill in the holes left by the removal 

of common buckthorn with soil and re-vegetate with native 

species (Moriarty, 2005). 

Herbicide application: Herbicides should be applied in the early 

spring or late fall when most native vegetation is dormant to 

minimize any potential non-target effects. Applying herbicides 

to the basal bark will leave dead standing trees. These trees 

can be left standing, allowing native vegetation to grow around 

them, or they may be removed depending on the management 

planned for the area (Moriarty, 2005). Stems may also be cut 

down and removed, but in this case herbicides need to be applied 

to the stump to prevent re-sprouting. This method may be time 

consuming and labour intensive because common buckthorn 

can have numerous stems and grow in dense thickets (Delanoy 

& Archibald, 2007). 




Areas with large thickets of common buckthorn will require 

time, effort and money to manage. To help reduce initial costs 

and ensure practicality, only female plants should initially be 

targeted for control. This will prevent seed production as male 

plants do not bear fruit. Preventing seed production helps to


control the populations and will help to decrease recruitment, 

thereby leading to a more manageable scenario. Control should 

occur in the fall to reduce detrimental effects on nesting 

birds. The dark coloured berries will be very prominent in the 

fall, making identification of female plants easier (Delanoy & 

Archibald, 2007). 

Although seed production is eliminated once all mature female 

plants are removed, seedlings may continue to emerge from 

an established seed bank. These seedlings may be pulled or 

treated with a basal herbicide treatment. When the invasion has 

become somewhat manageable, male plants may be removed 

in a similar fashion to that used for the females. Seeding or 

planting native species may be required to restore the native 

plant community (Delanoy & Archibald, 2007). 

Integrated control should be implemented when facing dense 

patches of common buckthorn in areas with desirable vegetation. 

Hand-pulling small saplings decreases the amount of herbicide 

required. However, herbicides will be needed to kill larger trees 

that cannot be pulled or dug out from the ground without major 

soil disturbance or mechanical help. Alternatively, herbicides 

may be applied to the freshly cut stump to prevent any shoots 

from sprouting and to eliminate the need to dig out a large root 

system (Delanoy & Archibald, 2007). 


Managing large common buckthorn invasions without the use 

of herbicides will require a greater amount of time and effort. 

Target the large berry producing female trees to help control 

seed production (Delanoy & Archibald, 2007). These large trees 

and shrubs can be cut and left on site or placed in a brush pile for 

burning. The stumps will continue to re-sprout and will require 

frequent clipping or removal of the entire root system. Keep in 

mind that removing large stumps will cause soil disturbance 

that could promote reinvasion by common buckthorn or other 

invasive plants (Moriarty, 2005). 

After all the large common buckthorn trees and saplings have 

been removed, management efforts can focus on pulling the 

understory seedlings and saplings. Pulled plants may be placed 

in a slash pile for burning or left on site, ensuring that the roots 

are fully exposed to promote desiccation. Ongoing monitoring 

will be needed to control any seedlings emerging from a 

potential seed bank (Delanoy & Archibald, 2007). 

[ 141 ]

[ 142 ] 


Extent.of. 

infestation 

Table 13: Management recommendations for common buckthorn (Rhamnus cathartica). 

Recommended. 


Small.invasions.and.satellite.populations Large.invasions.and.dense.populations 

Manual.and.mechanical.control:.handpulling.and.excavation. 

Integrated.control:.hand-pulling.and. 

herbicide.application. 

Timing Spring,.summer.and.fall. Fall.herbicide.application..Hand-pulling. 

can.occur.in.spring,.summer.or.fall. 

Disposal Woody.debris.may.be.left.on.site.or. 

removed.to.a.brush.pile.or.burned. 

Frequency.of. 

control 

Initial.pull.should.remove.as.many.trees. 

and.shrubs.as.possible..Return.several. 

times.per.year.to.pull.any.missed.plants.or. 

new.sprouts. 


Required. 

restoration 


hand-pulling.and.excavating.creates.soil. 

disturbance. 

Dead.shrubs.and.woody.debris.may.be. 

left.on.site.or.removed.to.a.brush.pile. 

or.burned. 

Initial.pull.should.remove.as.many.trees. 

and.shrubs.as.possible..Return.several. 

times.per.year.to.pull.any.missed.plants. 

or.new.sprouts..Use.herbicidal.control. 

once.every.fall. 


hand-pulling.and.excavating.creates. 

soil.disturbance.


5.1.9 

Periwinkle 

(Vinca minor) 


common periwinkle, lesser periwinkle, Myrtle, running myrtle 


143 

Figure 158: Periwinkle [ ] 

1 [ ]

[ 144 ] 



steM: Periwinkle is a trailing vine growing as a carpet over 

the forest floor (Fig. 159). The stems contain a milky substance 

(Jenkins & Jackman, 1941) and can become slightly woody with 

age making them difficult to break by hand (Miller, 2003). 

leAves: Evergreen leaves are ovate in shape with a pointed 

tip (Fig. 160). These shiny, dark green leaves can be found in 

an opposite arrangement along the stem (Kaufman & Kaufman, 

2007) (Fig. 161). The leaf edges of periwinkle are smooth 

(Gleason & Cronquist, 1963). 

Figure 159: Periwinkle is a trailing 

vine 1 . 

Figure 160: Periwinkle leaves 1 . 

Flowers: Flowers have 5 petals and range in colour from 

blue to purple (Fig. 162). Solitary flowers are produced on short 

stems arising from leaf axils (Bailey, 1969). 

Fruit/seed: The fruit is a follicle that contains 3 to 5 black 

seeds. Seeds are produced occasionally however this type of 

reproduction is rare compared to vegetative spread (Stone, 

2009). 

Figure 162: Periwinkle flowers 1 . 

Figure 161: Opposite leaf 

arrangement 1 .

5.1.9 PERIWINKLE (Vinca minor) 


creeping snowberry (Gaultheria hispidula) 

Creeping snowberry is an evergreen plant with creeping stems. 

It can easily be distinguished from periwinkle by its alternate 

leaves with bristled edges. Periwinkle has opposite leaves with 

smooth edges (Fig. 163). Creeping snowberry leaves are also 

much smaller (

[ 146 ] 


common bearberry (Arctostaphylos uva-ursi) 

Common bearberry is similar to periwinkle in that it has 

trailing stems and evergreen leaves with smooth edges. It can 

be distinguished from periwinkle by its alternately arranged 

leaves (Dickinson et al. 2004) (Fig. 165). 

Figure 165: Comparing common bearberry 17 (left) and periwinkle 1 (right). 

twinflower (Linnaea borealis) 

Twinflower has creeping stems with opposite evergreen leaves 

similar to those of periwinkle. However, the leaves of twinflower 

are hairy with toothed edges while the leaves of periwinkle are 

hairless with smooth edges (Dickinson et al. 2004) (Fig. 166). 

Figure 166: Comparing twinflower 20 (left) and periwinkle 1 (right).


winter creeper (Euonymus fortunei) 

Winter creeper shares several similarities with periwinkle. Its 

opposite evergreen leaves and trailing habit are similar to those 

of periwinkle as is its invasive nature in the forest understory. 

It is native to Japan, China and Korea. Winter creeper has 

leaves with toothed edges as opposed to the smooth edges of 

periwinkle leaves (Kaufman & Kaufman, 2007) (Fig. 167). 

Figure 167: Comparing winter creeper 12 (left) and periwinkle 1 (right). 

Partridge-berry (Mitchella repens) 

Partridge-berry can be easily confused with periwinkle. A 

trailing habit, opposite evergreen leaves with smooth edges, and 

light green veins are all characteristics that these two species 

share. The easiest way to identify these species is to look at the 

leaf tip. Partridge-berry leaves have a very rounded tip whereas 

the leaves of periwinkle have pointed tips (Chambers et al. 

1996) (Fig. 168). 

Figure 168: Comparing partridge-berry 1 (left) and periwinkle 1 (right). 

[ 147 ]

[ 148 ] 


A key to evergreen plants that may be confused 

with periwinkle (Vinca minor) 

1..Plants.with.alternate.leaves 

. 2..Leaves.with.bristled.edges 

. . 3..Small.leaves.(less.than.1cm.long)................................................Creeping.snowberry.(Gaultheria hispidula) 

. . 3..Large.leaves.(2-5cm.long)..............................................................Wintergreen.(Gaultheria procumbens) 

. 2..Leaves.with.smooth.edges................................................................Common.bearberry (Arctostaphylos uva-ursi) 

1..Plants.with.opposite.leaves 

. . . 4..Leaves.with.toothed.(serrate).edges 

. . . . 5..Leaves.covered.with.hair......................................................Twinflower.(Linnaea borealis) 

. . . . 5..Leaves.without.hair................................................................Winter.creeper.(Euonymus fortunei) 

. . . 4..Leaves.with.smooth.(entire).edges 

. . . . . 6..Leaves.with.rounded.tips................................................Partridge-berry.(Mitchella repens) 

. . . . . 6..Leaves.with.pointed.tips.................................................Common.periwinkle.(Vinca minor) 






........Subclass. Asteridae 

..........Order. Gentianales 

............Family. Apocynaceae 

BioloGy 


Periwinkle was introduced by European 

settlers in the early 1700’s (Swearingen 

et al. 2010). A native to Eurasia, it was 

valued as an easy-to-grow ornamental 

groundcover (Winterrowd & Stagg, 

1993). Today it is still commonly sold 

as a garden plant throughout the 

province (Miller, 2003). 

hABitAt 

..............Genus. Vinca 

Periwinkle is a versatile plant that can 

................Species. Vinca minor 

grow in open sunny areas to closed 

canopy forests (Swearingen et al. 2010). 

It is a common garden plant that has escaped cultivation to be 

found in open areas such as meadows and fields, edge habitats 

such as roadsides and along trails, and in forested areas (Fig. 

169). It is commonly seen growing as a dense cover in shaded 

areas (Miller, 2003; Stone, 2009).


Figure 169: Periwinkle growing alongside a road 1 (left) and trail 1 (right). 

reProduction 

Reproduction is mainly vegetative through stem runners. 

When these runners touch the soil at the leaf nodes they have 

the ability to take root (Jenkins & Jackman, 1941). Seeds are 

produced occasionally. However, this type of reproduction is 

rare compared to vegetative spread (Stone, 2009). These plants 

can easily establish from stem fragments (Winterrowd & Stagg, 

1993). 

liFe cycle 

Periwinkle is a perennial vine (Bailey, 1969) that flowers in 

the early spring (Gleason & Cronquist, 1963). The majority 

of flowers appear in April and May with occasional flowers 

appearing throughout the summer months. Seed-filled follicles 

are produced from May to July. Seeds are dispersed as the 

follicles dry out and crack. Periwinkles are evergreen and thus 

keep their leaves throughout the winter (Miller, 2003). 


Although the seed dispersing capabilities of periwinkle are 

low, these plants can propagate relatively quickly. Plants can 

send out twining stems in all directions that can take root 

wherever there is an empty space on the forest floor (Darcy 

& Burkart, 2002). Since these plants are free-rooting, even 

small stem fragments have the ability to grow into a dense mat 

(Fig. 170). Such plant fragments can become established 

throughout the spring and summer months (Winterrowd & 

Stagg, 1993). 

Periwinkle can effectively compete with native vegetation in 

the forest understory. This competitive advantage may be due 

to allelopathy, light suppression or a combination of both. A 

[ 149 ]

[ 150 ] 


laboratory study suggested that periwinkle had allelopathic 

effects on the growth rate of tree seedlings. However, more 

studies are needed to fully understand these effects (Darcy & 

Burkart, 2002). 

Periwinkle is an evergreen vine that has the ability to grow 

over other plants. This growth habit gives the plant the best 

access to sunlight. The native plants growing underneath 

are at a disadvantage as they cannot get enough sunlight for 

photosynthesis (Darcy & Burkart, 2002). 

Figure 170: Periwinkle forming a carpet on the forest floor 1 . 


Due to periwinkle’s limited dispersal capabilities it is not 

always considered a major threat. However, once established 

this species may have negative effects on the woodlot. Control 

is often difficult due to the free-rooting nature of this 

species. Monitoring the woodlot for invasions by periwinkle is 

recommended. 

Periwinkle has the ability to suppress hardwood regeneration. 

It forms a dense carpet (Fig. 171) that covers low growing 

species (Drake et al. 2003). Through light suppression and the 

possibility of allelopathic effects, seedling growth is inhibited. 

With the absence of regeneration, the species composition of 

the forest may gradually change (Darcy & Burkart, 2002). 

Periwinkle may also have negative impacts on the understory 

fauna. One study showed a change in the spider community 

associated with the forest floor. Such changes have a ripple


effect where impacts may be felt throughout the local food web 

(Bultman & DeWitt, 2008). More studies are needed to document 

such underlying effects. 

Figure 171: Periwinkle growing as a carpet on the forest floor 1 . 


Humans are the main vectors of periwinkle spread. Periwinkle 

is still a common garden plant and it is sold in garden centres 

throughout Ontario (Fig. 172). It is unlikely that seeds are a 

dispersal agent as these plants rarely germinate from seed 

in North America. Most spread is due to the dumping of yard 

wastes in natural areas. These plants can easily propagate from 

the smallest root fragments, thus such yard wastes are a major 

factor in the spread of this species (Stone, 2009). 

Figure 172: Periwinkle being sold as a ground cover 1 . 

[ 151 ]

[ 152 ] 




• Do not dump yard wastes in or close to natural areas; 

• Periwinkle is considered an invasive plant and should not be 

planted in the garden. Use native alternatives or species that 

are not considered invasive. 


• Learn how to identify periwinkle; 

• Regularly monitor the woodlot; 

• Become familiar with periwinkle populations in neighbours 

gardens and ensure that yard wastes are properly disposed of. 


Hand-pulling: Hand-pulling can be labour intensive and is only 

suitable for small invasions or for outlying satellite populations. 

All plant fragments need to be collected and bagged as they 

can easily propagate. Areas where runners have rooted into 

the ground need to be pulled. Using a rake will help to collect 

the twining stems while raising the runners to reveal the root 

system. Digging tools may be required to ensure removal of the 

entire root system since any portion of the root left behind has 

the ability to re-sprout. As the stems can be slightly woody it 

is recommended to have pruning shears on hand (Swearingen 

et al. 2010). 

Herbicide application: Chemical control is suitable for severe 

invasions. Herbicides should be applied as a foliar spray, 

enough to wet leaves but not so much as to cause dripping. 

The leaves of periwinkle plants have a waxy cuticle that 

does not readily absorb herbicides (Bean & Russo, 1988). The 

addition of a surfactant to the herbicide solution will increase 

absorption by the leaves and increase the likelihood of success.


Repeated treatments and spot applications are often necessary. 

Alternatively, the cut-and-spray method may be used to increase 

the chance of absorption through the wounded plant tissues 

(Drewitz, 2000). 




The use of chemical control is recommended during the fall 

when most native plants are dormant. The most effective 

means of control may be the cut-and-spray method. This 

method combines both physical and chemical control. The 

stems are cut and herbicides applied immediately afterwards. 

The wounded stems will readily absorb the herbicide solution 

(Drewitz, 2000). Herbicides may also be applied as a foliar 

spray. A second application may be needed the following year 

if the initial application was unsuccessful. Periwinkle plants 

that are in close proximity to desirable vegetation may be handpulled 

to prevent any damage to native species. It is also good 

to keep in mind that a combination of manual and chemical 

control decreases the amount of herbicides entering the forest 

ecosystem while managing cost and labour requirements (Bean 

& Russo, 1988). 

option # 2: without chemical control 

Hand-pulling can have a minimal impact on native vegetation 

if care is taken not to trample any native species. However, 

frequent monitoring is required and it has the potential to be 

labour intensive. The initial pull should be focused on removing 

as many plants and plant fragments as possible. Bag all parts of 

the plant and dispose of them at an appropriate landfill. Return 

to the area several times per year to pull any missed plants or 

new sprouts. Hand-pulling often creates soil disturbance which 

is known to increase the likelihood of reinvasion. It may be 

beneficial to plant desirable native seedlings in areas where 

such soil disturbing activities have occurred (Swearingen et al. 

2010). 

[ 153 ]

[ 154 ] 


Table 14: Management recommendations for periwinkle (Vinca minor). 


Recommended. 


Manual.control:.hand-pulling. Integrated.control:.cutting,.handpulling.and.herbicide.application. 

Timing Spring,.summer.and.fall. Fall. 

Disposal Bag.all.parts.of.the.plant.and.dispose.at. 

an.appropriate.landfill. 

Frequency.of.control Initial.pull.should.remove.as.many. 

plants.and.plant.fragments.as.possible.. 

Return.several.times.per.year.to.pull.any. 

missed.plants.or.new.sprouts. 


Required.restoration Plant.native.species.in.areas.where. 

hand-pulling.creates.soil.disturbance. 

Plant.material.may.be.left.on.site. 

Once.yearly.until.invasion.has.been. 

eliminated.or.has.become.small.enough. 

to.manage.with.manual.control. 


hand-pulling.creates.soil.disturbance.


5.1.10 

dog-strangling vine 

(Vincetoxicum rossicum & V. nigrum) 


european swallow-wort, Pale swallow wort, louis’ swallow-wort, 

Black swallow-wort 


155 

Figure 173: Dog-strangling vine [ ] 

1 [ ]

[ 156 ] 



There are two species of dog-strangling vine currently invading 

Ontario habitats. They are Vincetoxicum rossicum and 

V. nigrum. A third species, V. hirundinaria has not yet invaded 

North America and thus will not be discussed herein. However, 

it could potentially become a problem in the future (see 

extensive review by DiTommaso et al. 2005). 

leAves: Both species have similar leaves that are ovate with 

smooth edges (Fig. 174). They occur in an opposite arrangement 

along the stem (DiTommaso et al. 2005). 

Figure 174: Dog-strangling vine leaves 1 . 

Flowers: Both species of dog-strangling vine have flowers 

with 5 petals. Vincetoxicum rossicum flowers range in colour 

from pink to dark maroon (Fig. 175). They form clusters of 5 to 

20 flowers that sprout from leaf axils (Fig. 176). Vincetoxicum 

nigrum flowers can be dark purple to almost black in colour 

and form clusters of 4 to 10 flowers (DiTommaso et al. 2005). 

Figure 175: Comparison of flower colours for Vincetoxicum rossicum 1 

(left) and V. nigrum 6 (right). 

Figure 176: Vincetoxicum rossicum 

flower clusters 1 .

5.1.10 DOG-STRANGLING VINE (Vincetoxicum rossicum & V. nigrum) 

Fruit/seed: Generally, two fruits in the form of follicles are 

produced per V. rossicum flower (Fig. 177) whereas V. nigrum 

flowers will only rarely produce two fruits per flower. Seeds 

are connected to a tuft of white hairs (Fig. 178) and are similar 

to those of native milkweed (DiTommaso et al. 2005). 

Figure 177: Vincetoxicum rossicum follicles 1 . Figure 178: Dog-strangling vine seed dispersal 6 . 


Amur honeysuckle (Lonicera maackii) 

Amur honeysuckle is a shrub that may be confused with young 

dog-strangling vine plants that have not yet started to climb or 

look like a vine. They both have opposite, oblong leaves with 

smooth edges (Fig. 179). Amur honeysuckle leaves have long 

pointed tips whereas dog-strangling vine has leaves with a 

much shorter tip (Kaufman & Kaufman, 2007). 

Figure 179: Comparing Amur honeysuckle 1 (left) and dog-strangling 

vine 1 (right). 

[ 157 ]

[ 158 ] 


Morrow’s honeysuckle (Lonicera morrowii) 

The shape and edges of a morrow’s honeysuckle leaf is very 

similar to that of a dog-strangling vine leaf (Fig. 180). However, 

one key feature of morrow’s honeysuckle is that the underside 

of each leaf is covered in hair, whereas the leaves of dogstrangling 

vine are hairless (Kaufman & Kaufman, 2007). 

Figure 180: Comparing morrow’s honeysuckle 1 (left) and dog-strangling vine 1 (right). 

canada fly honeysuckle (Lonicera canadensis) 

Another shrub with similar looking leaves to dog-strangling 

vine is the Canada fly honeysuckle (Fig. 181). The leaf edges of 

Canada fly honeysuckle are ciliate which means that they are 

fringed with tiny hairs (Fig. 182). These hairs can be seen on 

close inspection with a magnifying glass. The leaves of dogstrangling 

vine are hairless (Chambers et al. 1996). 

Figure 181: Comparing Canada fly honeysuckle 1 (left) and dogstrangling 

vine 1 (right). 

tartarian honeysuckle (Lonicera tatarica) 

Tartarian honeysuckle leaves are similar to those of dogstrangling 

vine in that they do not have any hair on their 

surface or edges. However, some of the older leaves on tartarian 

honeysuckle have a heart-shaped appearance at the base (Fig. 

183). This is usually not seen on leaves found close to the end 

Figure 182: Ciliate leaves of Canada 

fly honeysuckle 1 .


of the twigs so be sure to check farther down the stem (Kaufman 

& Kaufman, 2007). 

Figure 183: Comparing the leaves of tartarian honeysuckle 1 (left) and dog-strangling vine 1 (right). 

Bush honeysuckle (Diervilla lonicera) 

On close inspection bush honeysuckle’s leaves can easily be 

distinguished from those of dog-strangling vine by the appearance 

of the leaf margin (Fig. 184). The leaves of bush honeysuckle have 

a toothed margin whereas the leaves of dog-strangling vine are 

smooth along the edges (Chambers et al. 1996). 

Figure 184: Comparing bush honeysuckle 20 (left) and dog-strangling vine 1 (right). 

dogwood (Cornus spp.) 

Some dogwood species in Ontario may be confused with dogstrangling 

vine. Although dogwood species in Ontario are trees 

and shrubs, when young they may be mistaken for dog-strangling 

vine. The key to distinguish them is to look at the leaf veins 

(Fig. 185). The leaf veins of dogwood species curve to meet the 

leaf tip whereas dog-strangling vine leaves have veins that curve 

towards the outer margin (Chambers et al. 1996). 

Figure 185: Comparing the leaves of dogwood 1 (left) and dog-strangling vine 1 (right). 

[ 159 ]

[ 160] 


oriental & American bittersweet (Celastrus 

orbiculatus & C. scandens) 

Oriental and American bittersweet are vines with alternate 

leaves. The leaves of both species have toothed margins 

whereas dog-strangling vine has smooth margins (Fig. 186). 

Oriental bittersweet leaves are nearly round with a blunt tip 

while American bittersweet has oblong leaves with pointed tips 

(Kaufman & Kaufman, 2007). 

Figure 186: Comparing Oriental bittersweet 15 (left) and dog-strangling vine 1 (right). 

Bittersweet nightshade (Solanum dulcamara) 

Bittersweet nightshade is a climbing vine with alternate leaves 

and smooth margins. Many of the leaves have two leaflets at the 

base (Fig. 187). The flowers and fruit of bittersweet nightshade 

are dissimilar to those of dog-strangling vine (Fig. 188). 

Bittersweet nightshade has seeds encased in berries whereas 

dog-strangling vine has the characteristic seedpods belonging 

to the milkweed family (Dickinson et al. 2004). 

Figure 187: Comparing the leaves of bittersweet nightshade 1 (left) to those of dog-strangling vine 1 (right).


Figure 188: Comparing the flowers of bittersweet nightshade 1 (left) to those of dog-strangling vine 1 (right). 

honeysuckles (Lonicera spp.) with climbing stems 

Honeysuckles with a climbing habit can be very similar to dogstrangling 

vine. They all have an opposite leaf arrangement. 

Three honeysuckle species, hairy honeysuckle (Lonicera 

hirsuta), smooth honeysuckle (L. dioica) and coral honeysuckle 

(L. sempervirens), can be excluded by looking at the leaf pairs 

located at the end of the vine (Fig. 189). These leaf pairs are 

joined together at the base so that the two leaves appear to be a 

single leaf that encircles the stem (Chambers et al. 1996). 

Japanese honeysuckle (L. japonica) is very similar to dogstrangling 

vine. It may be useful to look for a lighter coloured 

green on the bottom surface of its leaves (Fig. 190). However, it 

may be better to check for the milky sap from crushed leaves 

and stem characteristic of dog-strangling vine when trying to 

distinguish these species (Kaufman & Kaufman, 2007). 

Figure 189: The fused leaves of trumpet honeysuckle 16 . Figure 190: Japanese honeysuckle leaves 12 . 

[ 161]

[ 162] 



dog-strangling vine (Vincetoxicum spp.) 

1..Plants.with.a.woody.stem.(shrub).and.opposite.leaves.(for.specimens.where.stems.may.not.seem.woody) 

. 2..Leaves.with.sharply.pointed.edges.(toothed.margins).......................Bush.honeysuckle.(Diervilla lonicera) 

. 2..Leaves.with.smooth.edges.(entire.margins) 

. . 3..Leaves.with.narrow,.long-pointed.tips................................................Amur.honeysuckle.(Lonicera maackii) 

. . 3..Leaves.with.short,.blunt.tips 

. . . 4..Leaves.with.hair.on.surface.or.edges.(margins) 

. . . . 5..Underside.of.leaf.covered.in.fine.hairs........................................Morrow’s.honeysuckle.(Lonicera morrowii) 

. . . . 5..Underside.of.leaf.hairless,.edges.ciliate......................................Canada.fly.honeysuckle.(Lonicera canadensis) 

. . . 4..Leaves.with.no.hair.on.surface.or.edges.(margins) 

. . . . . 6..Leaf.veins.curve.to.meet.tip.of.leaf.........................................Dogwood.(Cornus.spp.) 

. . . . . 6..Leaf.veins.curve.towards.margins. 

. . . . . . 7..Leaves.slightly.heart-shaped.at.base..................................Tartarian.honeysuckle.(Lonicera tatarica) 

. . . . . . 7..Leaves.tapered.at.base............................................................Dog-strangling.vine.(Vincetoxicum.spp.) 

1..Plants.with.a.climbing.stem.(vine) 

. . . . . . . 8..Alternate.leaves 

. . . . . . . . 9..Leaves.with.smooth.edges.(entire.margins)...........Bittersweet.nightshade.(Solanum dulcamara) 

. . . . . . . . 9..Leaves.with.sharply.pointed.edges.(toothed.margins) 

. . . . . . . . . 10..Leaves.nearly.round.with.short.tip.....................Oriental.bittersweet.(Celastrus orbiculatus) 

. . . . . . . . . 10..Leaves.oblong.with.pointed.tip...........................American.bittersweet.(Celastrus scandens) 

. . . . . . . 8..Opposite.leaves 

. . . . . . . . . . 11..Upper.leaf.pairs.joined.together.at.base 

. . . . . . . . . . . 12..Leaves.with.hair.on.surfaces.........................Hairy.honeysuckle.(Lonicera hirsuta) 

. . . . . . . . . . . 12..Leaves.with.no.hair..........................................Smooth.honeysuckle.(Lonicera dioica).& 

. . . . . . . . . . . . ................................................................................Coral.honeysuckle.(Lonicera sempervirens) 

. . . . . . . . . . 11..Upper.leaf.pairs.distinct.(not.joined) 

. . . . . . . . . . . . 13..Underside.of.leaf.a.paler.green.colour...Japanese.honeysuckle.(Lonicera japonica) 

. . . . . . . . . . . . 13..Both.leaf.surfaces.similar.in.colour..........Dog-strangling.vine.(Vincetoxicum spp.)


Dog-strangling vine.(Vincetoxicum rossicum) 







............Family. Asclepiadaceae 

..............Genus. Vincetoxicum. 


................Species. Vincetoxicum rossicum. 

BioloGy 


Vincetoxicum rossicum is native to the Ukraine and Russia 

while V. nigrum is native to southwestern Europe (Milbrath, 

2010). It is assumed that both species were introduced into 

North America as horticultural plants (DiTommaso et al. 2005). 

In Ontario, V. rossicum was first collected in Toronto in 1889 

(Smith et al. 2006). Today, it can be found invading areas from 

London to Ottawa. Vincetoxicum nigrum was often confused 

with V. rossicum and the date of its introduction into Ontario 

is unknown. Vincetoxicum nigrum has a wider distribution in 

Ontario than V. rossicum with populations scattered throughout 

southern and eastern Ontario (DiTommaso et al. 2005). 

hABitAt 

Dog-strangling vine.(Vincetoxicum nigrum) 







............Family. Asclepiadaceae 

..............Genus. Vincetoxicum 

................Species. Vincetoxicum nigrum 

Both species of dog-strangling vine favour areas exposed to full 

or partial sunlight (Smith et al. 2006) but will also grow in open 

forests (Kaufman & Kaufman, 2007). As with most invasive 

species, they will readily invade disturbed areas such as 

roadsides. Once established, long distance seed dispersal allows 

satellite populations to establish in natural areas (DiTommaso 

et al. 2005). Dog-strangling vine can thrive in a wide range of 

soils and differing light and temperature conditions (Smith et 

al. 2008; Sanderson & Antunes, in press). 

[ 163]

[ 164] 


reProduction 

Dog-strangling vines reproduce sexually and vegetatively 

through clones sprouting from an underground rhizome 

(Lumer & Yost, 1995). Plants self-pollinate if insect pollinators 

are unavailable for cross pollination (DiTommaso et al. 2005). 

Seeds are polyembryonic which means that each individual 

seed can give rise to a maximum of four seedlings (Ladd & 

Cappuccino, 2005). 

liFe cycle 

Dog-strangling vine is perennial. Flowers begin to emerge in the 

middle of May and last until late summer (Lumer & Yost, 1995). 

Green seedpods, similar to those produced by native milkweed, 

emerge at the end of June. These pods break open to release 

seeds from September through November. Dead brown vines 

remain entangled around supporting vegetation throughout 

the winter months (DiTommaso et al. 2005). 


With increasing size, dog-strangling vine populations can 

completely suppress native seedlings (Cappuccino, 2004). In low 

light environments, such as forest understories, dog-strangling 

vine has the ability to climb over native vegetation (Fig. 191). 

This allows the plant to obtain light while effectively reducing 

that available to the lower growing vegetation. As populations 

become larger they reduce space and nutrients available for 

other plants (Smith et al. 2006). 

The robust rootstalks of dog-strangling vine store water that 

may be used at times of drought. These rootstalks also provide 

an energy source that allows plants to grow early in the season 

giving them a head start over competing native vegetation 

(DiTommaso et al. 2005). 

Dog-strangling vine populations can increase rapidly due 

to their high reproductive potential (Fig. 192). Individual 

plants produce large quantities of seed. Even under shaded 

conditions where seed productivity is at its lowest, plants can


Figure 191: Dog-strangling vine in a forest understory 1 (left) and climbing over native vegetation 1 (right). 

produce as many as 28 000 seeds per square metre. Since 

seeds are polyembryonic, the number of seedlings produced 

may increase fourfold (Smith et al. 2006). Factors contributing 

to such a large seed production include the ability to selfpollinate 

and a long flowering season. Self-pollination ensures 

that each plant will produce seed (DiTommaso et al. 2005) 

while long-lived flowers allow for a greater fruit-set (Lumer & 

Yost, 1995). 

Figure 192: Numerous seedpods in a dog-strangling vine invasion 1 . 

[ 165]

[ 166] 


Another factor lending to this species success is the high 

survival rate of first-year plants. Due to polyembryony, each 

seed has more than one chance at occupying a site. Seedling 

emergence does not always occur at the same time. It may take 

up to 20 days before all seedlings have emerged from a single 

polyembryonic seed. As such, new seedlings can emerge to 

offset control practices (Ladd & Cappuccino, 2005). 

Dog-strangling vine has no known natural enemies in North 

America. Only minimal damage has been reported by insects 

or disease (Milbrath, 2010). As with other members of the 

milkweed family, dog-strangling vine contains cardenolides 

which are toxic if consumed in large quantities (DiTommaso et 

al. 2005). Thus, grazing wildlife such as deer are unlikely to eat 

these plants (Milbrath, 2010). 


Healthy regeneration is an important woodlot management 

goal. Dog-strangling vine can negatively impact forest 

regeneration by climbing over saplings (Fig. 193) and reducing 

access to available space, nutrients and light (DiTommaso et al. 

2005). Dog-strangling vine form symbiotic relationships with 

endomycorrhizal fungi and studies have shown that the number 

and types of fungi in soils are altered by large invasions. Such 

changes in soil biota can have negative consequences on native 

plants (Smith et al. 2008). 

Wildlife diversity is a value-added feature in a woodlot. For 

example, wildlife viewing often enhances the sugar bush 

experience for customers. Dog-strangling vine populations 

have been found to decrease the diversity of both insects 

and birds. Recently, this invasive species was associated to 

population decreases of the monarch butterfly, which is a 

species-at-risk. Monarch butterflies normally lay their eggs 

on native milkweed. Dog-strangling vine is a member of the 

milkweed family and may attract monarchs in areas where 

native milkweed is scarce. This is detrimental since monarch 

larvae are unable to fully develop on dog-strangling vine (Ladd 

& Cappuccino, 2005). Dog-strangling vine also displaces native 

milkweed, thereby decreasing available host plants needed for 

reproducing monarchs (DiTommaso et al. 2005).


Figure 193: Dog-strangling vine climbing over native vegetation 1 . 


Dog-strangling vine will likely enter a woodlot via wind-driven 

seed dispersal. Being part of the milkweed family, the seeds 

are similar to native milkweed, fringed with hairs that allow 

the seeds to be airborne on windy days (DiTommaso et al. 

2005). Although most seeds are known to fall within only a 

few metres of their parent, it only takes one seed to become an 

established plant that can grow with the help of clonal spread 

or self-pollination. Seeds may also get caught in the fur of 

animals, on clothing or tire treads (Smith et al. 2006). 

[ 167 ]

[ 168] 




• Prevent seeds from entering the woodlot by limiting access 

when possible; 

• Wash all shoes and tires before entering the woodlot; 

• Do not let domestic animals roam freely in late summer and 

fall when seed dispersal begins, especially if there are known 

dog-strangling vine populations in or around the woodlot; 

• Minimize soil disturbance during woodlot management 

activities; 

• Volunteer to help eliminate dog-strangling vine populations 

in properties nearby. 


• Learn how to identify dog-strangling vine; 

• Regularly monitor the woodlot, paying attention to any 

unknown vines. 


Cutting: Cutting can be used as a means to prevent seed 

production. Clipping the plants at ground level and removing 

the stem and leaves is best done in late June. This timing is 

important because the plants will have used up the majority 

of their resources to produce flowers. Thus clipped vines will 

not have enough energy left over to re-sprout and produce 

seeds before the winter. Although these plants will survive, it 

is beneficial to prevent any seed addition to the seed bank until 

more effective management strategies can be employed (Averill 

et al. 2008). 

Excavation: One way to manually control small populations of 

dog-strangling vine is to dig and excavate the whole root system. 

Do not try to pull the root up by hand as the stems easily break 

off, leaving root crowns and fragments in the soil that can 

readily re-sprout. Use spades or shovels to dig underneath the 

root system. Plants should be removed before they produce 

seed to prevent additions to the soil’s seed bank. Place all plant 

parts in a plastic bag for disposal in an appropriate landfill 

(DiTommaso et al. 2005). Excavating should only be used on very 

small populations as it can be labour intensive and causes soil 

disturbance. Restoration efforts will be required to fill in the 

holes where root systems were removed. Restoration should also 

include re-planting the area with native vegetation to prevent reinvasion 

of the disturbed area (Lawlor & Raynal, 2002).


Herbicide application: In large invasions where native vegetation 

has been compromised, it is best to apply herbicides as a foliar 

spray (Lawlor & Raynal, 2002). Be sure to add a surfactant to 

help the herbicide penetrate the waxy leaf surface (Douglass et 

al. 2009). Avoid applying herbicides as a foliar spray to vines 

that have become intertwined with young trees (Lawlor & 

Raynal, 2002). 




In areas with large invasions the use of chemical control is 

recommended. As plants succumb to the herbicide they can 

be replaced by a competitive native plant. Do not start replanting 

until the population is sufficiently under control, as 

spot treatments or excavation may be needed to completely 

eliminate dog-strangling vine from the area. Performing such 

control measures around newly planted vegetation may prove 

to be too labour intensive if done before adequate control is 

achieved (Lawlor & Raynal, 2002). 

A combination of manual and chemical control may be needed 

to control dense patches of dog-strangling vine intertwined 

with desirable vegetation. The first step to integrated control is 

an initial application of herbicide using a foliar spray for dense 

patches, cutting in areas where the vine is intertwined with 

desirable vegetation. Chemical control should be done in the 

spring at the bud stage to allow for manual control later in the 

season (Lawlor & Raynal, 2002). 


Dog-strangling vine invasions are particularly difficult to 

control. Hand-pulling is not effective as the stems easily breakoff, 

leaving behind a root system that will quickly re-sprout. 

Excavating the entire root system is often successful. However, 

this method can quickly become too labour intensive to be a 

feasible management strategy in areas with large invasions. The 

disturbance caused by digging also promotes the establishment 

of other invasive plants, often negating any positive effects 

of control. In areas where chemical control is not an option 

it is best to try and control the invasion instead of opting for 

complete eradication. Frequently cutting back the plants will 

prevent seed production and thus slow the spread of invasion. 

Planting competitive native species may also help control dogstrangling 

vine’s invasive potential (Averill et al. 2008; Lawlor 

& Raynal, 2002). 

[ 169]

[ 170 ] 


Table 15: Management recommendations for dog-strangling vine (Vincetoxicum spp). 


populations 

Recommended. 



Manual.Control:.excavation Integrated.Control:.herbicide.application. 

and.cutting. 

Timing May-early.June. Herbicide.application:.Spring.. 

Cutting:.Late.June. 

Disposal Place.all.plant.parts.in.a.plastic.bag.and. 

dispose.of.in.an.appropriate.landfill.. 

Ensure.no.roots.are.left.behind.to. 

resprout. 

Frequency.of. 

control 

Several.times.per.year..After.initial. 

excavation.return.to.collect.any. 

resprouting.plants.from.root.fragments. 

that.were.missed. 


Required. 

restoration 

Soil.disturbance.created.during. 

excavation.must.be.addressed..Fill. 

in.holes.and.replant.with.native. 

vegetation. 

Place.all.clipped.plants.in.a.plastic.bag.and. 

dispose.of.in.an.appropriate.landfill.. 

Herbicide.application.and.clipping.once. 

yearly.for.up.to.5.years.to.ensure.that. 

all.root.systems.have.been.killed.and.all. 

seeds.eliminated.from.the.seed.bank. 

May.require.some.re-planting.to.fill. 

in.gaps.where.dense.patches.of.dogstrangling.vine.eliminated.native. 

vegetation.


5.1.11 

Goutweed 

(Aegopodium podagraria) 


Bishop’s goutweed, Bishop’s weed, Ground elder, Goat’s-foot, 

snow-on-the-mountain 

Priority Rating: ModerAte 


Goutweed is native to Eurasia. It is a perennial plant with 

creeping stems. Compound leaves are in groups of 3, with 3 or 

fewer leaflets per group (Fig. 194). Leaflets often have irregular 

lobes. The leaves are found in an alternate arrangement along 

the stem (Kaufman & Kaufman, 2007). Escaped populations of 

goutweed generally have leaves that are of a solid green colour. 

Many cultivars have leaves with white borders around their 

paler green leaves. Both varieties may be seen in natural areas; 

however, the solid green colour is the most common. White 

flowers are produced in clusters on top of flowering stalks 

(Garske & Schimpf, 2005) (Fig. 195). 

Figure 194: Two colour varieties of goutweed including solid green 1 (left) and a lighter, multi-coloured 

version 6 (right). 

[ 171 ]

[ 172 ] 


Figure 196: Goutweed invasion in a 

forest understory 6 

environMentAl & 


Goutweed will usually establish around forest edges where it 

can access higher light levels. Once established the creeping 

stems can quickly spread through the forest understory (Fig. 

196). Its aggressive growth pattern often displaces native 

understory plants, which may inhibit hardwood regeneration 

(Garske & Schimpf, 2005). 

control 

Goutweed has an extensive rhizome that must be removed 

or severely damaged to prevent re-sprouting. Hand-pulling 

can be an effective method of control if both the above and 

belowground portions of the plant are entirely removed. 

Leaving small root fragments behind will often lead to a new 

invasion. Mowing in areas with dense stands can be effective if 

frequently repeated throughout the season. Root reserves must 

be completely depleted to prevent new invasions. Covering 

the invaded area with a black tarp will prevent re-sprouting 

by eliminating access to sunlight. Pay particular attention to 

the tarp edges and hand-pull any sprouting individuals (Fig. 

197). Eventually, root reserves will be depleted and the tarp 

can be removed. Applying herbicides as a foliar spray may 

kill goutweed but repeated applications are usually required 

(Garske & Schimpf, 2005; Kaufman & Kaufman, 2007). 

Figure 195: Goutweed flowers 6 . 

Figure 197: Goutweed control 1 .


5.1.12 

oriental Bittersweet 

(Celastrus orbiculatus) 


Asian bittersweet, japanese bittersweet, Asiatic bittersweet, 

round-leaved bittersweet 



Oriental bittersweet is native to Japan, China and Korea (Kaufman 

& Kaufman, 2007). It is a perennial woody vine. Leaves are round 

with toothed margins, ending in an abrupt tip. The leaves have 

an alternate arrangement along the stem. Oriental bittersweet 

is dioecious with separate male and female plants. Only female 

plants produce the distinctive berry-like fruits with 3 seed 

compartments (Fig. 198). These compartments are encased in a 

yellow capsule that opens as the fruit ripens (Swearingen, 2006). 

The exotic and the native bittersweet are often mistaken for one 

another, which makes control difficult. The leaves of American 

bittersweet (Celastrus scandens) are not as circular and the tips are 

not as abruptly pointed as in Oriental bittersweet leaves (Fig. 199). 

Figure 198: Oriental bittersweet fruits 6 

[ 173 ]

[ 174 ] 




As a vine, Oriental bittersweet can girdle trees and smother 

understory vegetation (Fig. 200). It can establish in a forest 

understory and remain relatively unobtrusive until a canopy 

gap appears. As light penetrates through the canopy, Oriental 

bittersweet will quickly grow and climb over native species 

to compete for available light (Greenberg et al. 2001). Oriental 

bittersweet competes with native American bittersweet for space 

and nutrients. As a result, American bittersweet is declining 

(Steward et al. 2003). Hybridization between the two species is 

also becoming a problem, as valuable genes may be lost. It is very 

important to ensure that American bittersweet is not damaged 

during management activities (Swearingen, 2006). 

control 

Hand-pulling is only practical in areas with a few scattered 

individuals. The entire plant, including the roots, should be 

removed to prevent re-sprouting. For large vines that have 

become tangled around valuable trees it is best to make two cuts 

in the stems so that the portion within reach is severed. The root 

system can either be removed to prevent re-sprouting or regular 

cuttings can be made throughout the season to exhaust the root 

reserves (Kaufman & Kaufman, 2007). A combination of manual 

and chemical control is recommended for large invasions. A 

licenced exterminator using the basal bark or cut-stem methods 

can apply herbicides. Repeated applications may be required to 

kill the vines. Monitor the area and cut any re-sprouts or perform 

a second herbicide treatment (Swearingen, 2006). 

Figure 199: Comparing the leaves of American bittersweet 

15 (left) to those of Oriental bittersweet 15 (right). 

Figure 200: Oriental bittersweet girdling a tree 6 

(left) and smothering native vegetation 35 (right).


5.1.13 

exotic Bush honeysuckle 

(Lonicera spp.) 


Amur honeysuckle, Morrow’s honeysuckle, tartarian honeysuckle, and 

Bell’s honeysuckle 


There are several invasive honeysuckles that can negatively 

impact a hardwood stand. They include amur honeysuckle 

(Lonicera maackii), morrow’s honeysuckle (L. morrowii), 

tartarian honeysuckle (L. tatarica), and bell’s honeysuckle (L. 

x bella). Invasive honeysuckles were introduced from China 

and Korea. All are shrubs with simple, opposite leaves and 

showy flowers (Fig. 201). These flowers are white or pink in the 

spring but slowly fade to yellow later in the season (Fig. 202). 

An abundance of bright red berries are produced in the fall 

(Kaufman & Kaufman, 2007) (Fig. 203). 

Figure 201: Exotic bush honeysuckle leaves 1 . 


Figure 202: Exotic bush honeysuckle flowers 1 

[ 175 ]

[ 176 ] 


Figure 203: Exotic bush 

honeysuckle berries 1 



Exotic bush honeysuckles produce leaves earlier than most 

native species in the spring. Early spring ephemerals evolved 

as a means of acquiring light before canopy closure. However, 

bush honeysuckles can out-compete these understory species 

by towering over them with even earlier emerging leaves. An 

aggressive growth rate and ability to form a dense shrub layer 

can exclude native understory species and affect hardwood 

regeneration (Kaufman & Kaufman, 2007) (Fig. 204). The bright 

red fruits are very attractive to birds, which leads to the avian 

dispersal of seeds. However, these fruits are nutrient-poor, 

which may have negative effects on migrating birds (Williams, 

2005). Nest predation rates may also be higher for songbirds 

nesting in invasive bush honeysuckles. A study by Schmidt and 

Whelan (1999) found that American robin nests experienced 

greater rates of predation due to being less protected in invasive 

bush honeysuckles than in native shrubs such as hawthorns. 

control 

Hand-pulling is only practical for small seedlings or saplings. For 

larger shrubs, mechanical levers such as root wrenches are needed to 

pry up the root system. Shrubs can be left onsite as long as their roots 

are exposed and do not touch the soil. This will allow the roots to 

dry out and eliminates any chance of re-sprouting. Shrubs may also 

be cut and the roots dug out with a shovel. This can be very labour 

intensive and causes a large degree of soil disturbance. Clipping 

re-sprouts may eventually exhaust the root reserves. However, this 

approach requires constant vigilance and is only practical when 

dealing with a few individuals. Invasive bush honeysuckles can 

be controlled using herbicides. A licenced exterminator can apply 

the chemicals using the basal bark or cut-stump method (Williams, 

2005). 

Figure 204: Exotic bush honeysuckle in a forest understory 12


5.1.14 

kudzu 

(Pueraria montana var. lobata) 


japanese arrowroot, nepalem 



Kudzu is a perennial vine with a semi-woody stem. The leaves are 

compound with 3 lobed leaflets that may also be entire (Fig. 205). 

The leaves have hairy margins and are arranged alternately along 

the stem. The flowers are pink to purple and grow in long clusters 

(Fig. 206). The seeds are encased in flat, hairy seedpods. Kudzu was 

introduced from Japan (Kaufman & Kaufman, 2007). 

Figure 205: Comparing lobed 36 (left) and entire kudzu leaflets 12 (right). 

[ 177 ]

[ 178 ] 


Figure 206: The flowers 37 (left) and seedpods 36 (right) of kudzu. 



Kudzu grows in large mats, using other plants as supporting 

structures. The dense foliage often kills native vegetation by 

reducing light levels. The vining growth strategy girdles trunks 

and stems and the rapid and prolific growth rate creates large 

amounts of biomass that can crush supporting plants and uproot 

trees. Kudzu grows along forest edges but once established 

might spread into the forest interior as it destroys native edge 

species (Bergmann & Swearingen, 2005) (Fig. 207). 

Figure 207: Kudzu invading a forest edge 38 .


control 

Kudzu has an extensive and robust root system. Taproots 

can grow very large and weigh hundreds of pounds. The key 

to controlling an invasion of kudzu is to destroy the taproot 

through the exhaustion of nutrient reserves or through 

herbicide application (Kaufman & Kaufman, 2007). Large 

invasions should be cut and removed. However, since it is often 

impractical to remove the large taproots, repeated cutting is 

recommended. After this, close monitoring is required so that 

any new growth is immediately cut to prevent photosynthesis 

and prompt the roots to utilize energy to create new sprouts 

(Bergmann & Swearingen, 2005). 

Herbicide application is another method of controlling kudzu 

invasions. Herbicides are best applied using the cut-stump 

method. Since kudzu stands are usually dense the stems 

must be cut so that the herbicide can reach the interior of the 

stand. Herbicides should be applied by a licenced exterminator 

immediately after the stems have been cut. Monitor the area 

and clip any re-sprouts or perform a second application of 

herbicides as required (Bergmann & Swearingen, 2005). 

[ 179 ]

[ 180] 


5.2 

exotic insects & disease 

5.2.1 Emerald Ash Borer (Agrilus planipennis) HIGH 

5.2.2 Asian Long-horned Beetle (Anoplophora glabripennis) HIGH 

5.2.3 Chestnut Blight (Cryphonectria parasitica) HIGH 

5.2.4 Beech Bark Disease (Cryptococcus fagisuga & Neonectria spp.) HIGH 

5.2.5 Gypsy Moth (Lymantria dispar) HIGH 

5.2.6 Dutch Elm Disease (Ophiostoma spp.) HIGH 

5.2.7 Elm Bark Beetles (Scolytus multistriatus & S. schevyrewi) HIGH 

5.2.8 Butternut Canker (Ophiognomonia clavigignenti-juglandacearum) HIGH 

5.2.9 Dogwood Anthracnose (Discula destructiva) MODERATE 

5.2.10 Thousand Cankers Disease (Geosmithia morbida) MODERATE 

5.2.11 Pear Thrips (Taeniothrips inconsequens) MODERATE


5.2.1 

emerald Ash Borer 



eAB 


Figure 208: Emerald ash borer 39 . 

5.2 

181 

[ ]

[ 182] 


orGAnisM 


As an adult insect, the emerald ash borer (EAB) has a long, 

narrow body that ranges in length from 0.75 to 1.5cm. They are 

shiny with emerald green and metallic copper coloured hues 

(Fig. 209). Their black or copper eyes appear very large on their 

heads. The top of the abdomen is a bright metallic red colour 

that can be seen when the beetle is flying (Lyons et al. 2007) 

(Fig. 210). 

Figure 209: Adult emerald ash borer 39 . Figure 210: Characteristic metallic red abdomen of 

the emerald ash borer 39 . 

host trees 

The only species directly affected by EAB belong to the ash genus 

(Fraxinus spp.) (Kimoto & Duthie-Holt, 2006) and even exotic 

ash trees are vulnerable to EAB (Lyons et al. 2007). Emerald ash 

borer insects are primarily attracted to 

stressed ash trees, however, in heavy 

infestations even healthy individuals are 

attacked (Poland & McCullough, 2006). 

Ash trees are characterized by their 

oppositely arranged, pinnately divided 

compound leaves consisting of 5 to 11 

leaflets (Farrar, 1995)(Fig. 211). There 

are 16 species of ash in North America 

with 5 found in Ontario. The blue ash 

(F. quadrangulata) and pumpkin ash 

(F. profunda) are relatively rare, found 

mainly in southwestern areas of Ontario. 

Black ash (F. nigra) is common throughout 

Ontario, found mostly in wet, swampy 

areas. White ash (F. americana) and red 

ash (F. pennsylvanica) are common and 

economically valuable lumber species in 

Ontario (Kershaw, 2001). 

Figure 211: Compound leaf of black ash 

(Fraxinus nigra) 8 .

exit holes 

5.2.1 EMERALD ASH BORER (Agrilus planipennis) 

siGns & syMPtoMs 

The emerald ash borer is host specific to 

species of the genus Fraxinus; all ash trees 

in North America are susceptible. After 

larval development is complete, adult 

beetles emerge from their host tree creating 

D-shaped exit holes (Fig. 212). These holes 

are approximately 3.5mm wide and 4mm 

long. Exit holes can be found on the trunk 

and branches of ash trees (Kimoto & Duthie- 

Holt, 2006). 

FeedinG GAlleries & BArk dAMAGe 

Larvae feeding on the inner sapwood create long curving 

tunnels called feeding galleries (Fig. 213). These S-shaped 

feeding galleries can cause vertical cracks on the trunk (Fig. 

214). Galleries are generally 6mm wide and vary from 9 to 16cm 

long. On occasion, up to 30cm long galleries have been observed 

(Kimoto & Duthie-Holt, 2006). 

Figure 213: Feeding galleries 41 created by emerald 

ash borer larvae 

Adult FeedinG 

Emerald ash borer beetles feed on ash 

leaves immediately after emerging from the 

tree trunk as adults. Adult feeding creates 

notches along the edges of the leaves (Lyons 

et al. 2007) (Fig. 215). Several leaf notches 

may be seen along the outer margin. Feeding 

adults are most active during the day. They 

may be found seeking shelter under the 

leaves or in bark fissures during cold or wet 

weather (McCullough et al. 2008). 

Figure 212: D-shaped exit hole 

created by the emerald ash borer 40 . 

Figure 214: Bark damage caused by 

emerald ash borer feeding galleries 40 . 

Figure 215: Emerald ash borer 

feeding on an ash leaf 42 . 

[ 183]

[ 184] 


BrAnch die-BAck & ePicorMic shoots 

Feeding tunnels created by EAB larvae prevent the transport of 

sap in the tree, which usually results in death within 2 years. The 

top branches are usually the first to wilt. In response, ash trees 

produce new shoots along their lower trunk (Fig. 216), which can 

be used to detect EAB infestation (OMNR, 2010b). 

Figure 216: Crown dieback and epicormic shoots on an infested ash tree 43 . 

siMilAr sPecies, siGns & 

syMPtoMs 

other Agrilus spp. 

Several other beetles belonging to the Agrilus genus are found 

in Ontario. Most appear very similar to EAB. Some examples, 

including the bronze birch borer (A. anxius) and the metallic 

wood-boring beetle (A. cyanescens) are shown in Figure 217. 

Although very similar in appearance, these species do not 

reproduce on ash trees. The only other species of Agrilus known 

to attack ash in Ontario is A. subcinctus; however this species is 

much smaller than EAB, has a black body and the abdomen has 

a copper shine. Always look for additional signs and symptoms 

to help identify potential EAB infestations (GISD, 2006). 

Figure 217: Comparing A. anxius 44 (left) and A. cyanescens 44 (right).

Ash yellows 


The majority of ash species are susceptible to a disease called 

ash yellows. This disease is caused by a specialized group of 

bacteria (Candidatus Phytoplasma fraxini) that affects the 

phloem of the tree (Griffiths et al. 1999). The disease is spread 

by leafhoppers. Small sections of the crown will experience 

branch dieback as a result of phloem disruption (Fig. 218). In 

contrast, EAB infestations can cause branch dieback throughout 

the crown, eventually killing the tree (Sinclair & Griffiths, 1994). 

Figure 218: Ash showing symptoms of ash yellows disease 45 . 

Feeding Galleries & exit holes 

Various insect species can create similar tunnels and exit holes 

in ash trees that can be confused with those of EAB. Although 

the actual insects look nothing like EAB, their damage may 

appear similar. One example is the native red-headed ash 

borer (Neoclytus acuminatus). This red and yellow insect is 

quite distinct and its larvae feed on the sapwood of ash trees, 

creating feeding tunnels that can kill the tree. These tunnels 

are generally not as meandering as EAB’s S-shaped galleries 

(Fig. 219). The exit holes are circular unlike the distinctive 

D-shaped exit holes made by EAB (Herms, 2007). 

Figure 219: Red-headed ash borer, exit holes 46 and feeding galleries 18 . 

[ 185]

[ 186] 



BioloGy 


Kingdom. Animalia 

EAB is a bark beetle native to Asia, where 

it can be found in China, Japan, Korea, 

..Phylum. Arthropoda 

Mongolia, Taiwan and Russia (Haack 

....Class. Insecta 

et al. 2002). It was first discovered in 

North America in 2002 in Michigan, 

......Order. Coleoptera 

and shortly thereafter in Ontario. 

........Family. Buprestidae 

However, evidence indicates that it 

went unnoticed since the early 1990’s 

..........Genus. Agrilus 

(Siegert et al. 2007). EAB is very difficult 

............Species. Agrilus planipennis 

to detect, especially at the early stages 

of infestation. Any noticeable signs and 

symptoms are often attributed to other 

environmental stressors such as other insects, pathogens and 

weather events (Lyons, 2010). Since its detection in Michigan 

and Ontario in 2002, surveys have confirmed its presence in 

other states such as Indiana and Ohio (Cartwell, 2007). 

liFe cycle 

The emerald ash borer has a four-stage life cycle including 

egg, larva, pupa and adult (Fig. 220). Adult females deposit 

their eggs on the trunk or branches of ash trees during the 

summer months. Each individual female can produce 50 to 

90 eggs during their 3 to 6 week lifespan. Eggs hatch within 

2 weeks and the larvae bore into the sapwood creating S-shaped 

feeding galleries. Larvae slowly develop through 4 instars and 

stop feeding by November. These pre-pupae overwinter within 

the sapwood. Pupation begins in the spring and lasts for about 

3 weeks. After pupation, adult beetles create a D-shaped hole 

through which they emerge (Kimoto & Duthie-Holt, 2006; 

Poland & McCullough, 2006). 

Egg Larva 

A d u l t P u p a 

Figure 220: Life cycle of the emerald ash borer: egg 28 , larva 39 , pupa 39 and adult 43 .


iMPActs 

EAB is destroying ash populations in Ontario. The larvae 

feeding on the inner sapwood of ash trees effectively cut off 

the transportation of sap within the tree (Fig. 221). Mortality 

usually occurs within 2 to 4 years after EAB establishes in 

large mature trees but may only take 1 year for small trees 

and saplings. The 5 species of ash in Ontario contribute to 

forest biodiversity and are a source of food and habitat for 

wildlife (Poland & McCullough, 2006). Ash is also a popular 

ornamental tree in urban areas and is an important commercial 

lumber and pulp species. Considering that ash mortality due 

to EAB is 100%, the environmental, aesthetic and economic 

impacts caused by this invasive species represent a major 

challenge to Ontario (Cartwell, 2007). As natural regeneration 

of ash decreases due to a diminishing seed bank, the natural 

regeneration of ash in EAB-infested areas seems unlikely 

(Kashian & Witter, 2011). 

Figure 221: Larval tunneling damage causing tree mortality 39 . 

[ 187]

[ 188] 



The emerald ash borer was most likely introduced into North 

America through international trade in wood packaging 

material. Once in North America, it could easily be spread either 

by humans, through the transportation of wood products, or 

by flight during the adult life stage. Adult beetles have the 

potential to fly up to 7km per day (Taylor et al. 2010) and can 

create small colonies in previously uninfested areas, in some 

cases far removed from the main advancing front (Liebold & 

Tobin, 2008). However, humans are the leading cause of long 

distance dispersal, mainly through the movement of firewood 

(Petrice & Haack, 2007). 



Promoting a healthy woodlot while increasing forest diversity 

will help to deter EAB infestations. Healthy woodlots are often 

comprised of a wide range of species of different ages and sizes. 

This helps to decrease the number of available hosts, thereby 

creating poorer conditions for EAB invasion (OMNR, 2010b). 

In Ontario, a quarantine zone has been established by the CFIA, 

under the authority of the Plant Protection Act, to help control 

the spread of EAB. This quarantine restricts the movement of 

ash materials that could be vectors. These items include live or 

dead ash trees such as nursery stock, logs, leaves, lumber, wood 

and wood chips. In effect, it restricts the movement of any type 

of firewood (Poland & McCullough, 2006). This is important as 

studies have shown that EAB females will oviposit on ash trees 

after they have been cut (Anulewicz et al. 2008) and that EAB 

can successfully complete their life cycle on freshly cut material 

(Petrice & Haack, 2007). Indeed, preventive measures such as 

establishing quarantines do help to slow down the spread of 

EAB and other invasive forest pests (CFIA, 2011b). 


Visual surveys and monitoring activities are crucial for EAB 

detection. When walking through the woodlot it is best to take 

a close look at a few individual ash trees. Check for signs and 

symptoms of EAB invasion such as exit holes, epicormic shoots, 

and weak or dying ash trees. Ensure that monitoring activities 

are a regular routine. It is often difficult to detect EAB before 

the infestation becomes severe. This is because EAB spends


most of its life cycle underneath the bark (Cappaert et al. 

2005). Also, when EAB populations are low, the small number 

of larvae tunneling in the inner sapwood has little effect on 

halting sap transportation (McCullough et al. 2008). Emerald 

ash borer establishment on uninfected trees have frequently 

been observed to occur in the upper canopy as opposed to 

the trunk. This is one more reason why EAB infestations are 

difficult to detect (Anulewicz et al. 2007). 

Trap trees have been used as a means of detecting whether EAB 

is present in a woodlot. A trap tree is girdled with the intent 

of eliciting the release of stress related chemical signals that 

attract adult EAB (Poland & McCullough, 2010). Several studies 

have shown that EAB prefer weakened ash trees. However, they 

are equally capable of attacking healthy trees (McCullough et 

al. 2006; Poland et al. 2006). 

Figure 222: Emerald ash borer trap tree 40 . 

Constructed traps made with a sticky surface can also be used 

to trap EAB adults. These traps utilize visual and olfactory 

cues to attract beetles. Colour is an important visual cue for 

EAB. Purple and green are the most attractive colours likely 

because they mimic reflections of the trunk and leaves of ash 

trees as seen by EAB (Lyons, 2010). Olfactory cues are attractive 

to EAB because they mimic the odour of stressed ash trees 

(Rodriguez-Saona et al. 2006; de Groot et al. 2008; Crook et al. 

2008). Pheromones are also an olfactory cue that can be used as 

an attractant on the traps (Lyons, 2010). 

[ 189]

[ 190 ] 



Currently there are two ways of managing EAB infestations, 

including the removal of infested trees and the use of systemic 

insecticides. Infested trees can be cut down, chipped and burned 

(Fig. 223). Chipping and burning ensures that all life stages 

including the eggs, larvae, pupae and adults are destroyed. 

Trees should be cut down before the adults emerge in late May 

and June to prevent natural dispersal (Poland & McCullough, 

2010). Although this method can be both destructive and 

costly it should be done as early as possible. Infested trees will 

eventually die and may need to be removed anyway due to their 

potential to become a hazard to people. Moreover, removing 

and destroying infested trees helps to prevent further spread. 

Considering that EAB populations are already abundant 

and since some infested trees may show little or no sign of 

infestation, complete eradication is currently unattainable 

(Cartwell, 2007). 

Figure 223: Removing ash trees infested with emerald ash borer 43 . 

There is one systemic insecticide available for purchase in 

Canada that can be used in EAB control. TreeAzin was created 

by the Canadian Forest Service and BioForest Technologies 

Incorporated. This insecticide must be applied by a licenced 

exterminator and is only effective for a two-year period. 

Injecting ash trees with this insecticide will help to protect 

EAB free trees and may save some of those that are lightly 

infested (BioForest, 2012). Using TreeAzin for EAB control may 

be practical for use in woodlots with a very small percentage 

of ash trees or on ornamental trees in urban areas. Conversely, 

it can quickly become impractical in areas with large numbers 

of ash. The insecticide must be reapplied biannually in areas 

where EAB populations persist, which can be costly (Lyons, 

2010).


5.2.2 

Asian long-horned Beetle 

(Anoplophora glabripennis) 


Asian longhorn beetle, AlB, Basicosta white-spotted longicorn beetle, 

starry sky beetle 


Figure 224: Asian long-horned beetle 49 . 

191 

[ ]

[ 192 ] 


orGAnisM 


Adult Asian long-horned beetles are black with a glossy coating 

that reflects blue. They range from 2 to 3.5cm long. Adult 

females have antennae that are as long as their body whereas 

adult males have antennae that are twice their body length 

(Fig. 225). These antennae are made up of 11 segments with an 

alternating white and black colour. Approximately 20 white or 

yellow spots (Fig. 226) are found on each wing cover (Ric et al. 

2007). 

Figure 225: Comparing a female 50 (left) and a male 50 (right) Asian longhorned 

beetle. 

host trees 

The Asian long-horned beetle only attacks hardwood trees 

(i.e., deciduous trees). However, some hardwood species are 

more susceptible to the insect than others. They include maple 

(Acer sp.), birch (Betula sp.), poplar (Populus sp.), willow (Salix 

sp.), elm (Ulmus sp.), horsechestnut (Aesculus sp.), sycamore 

(Platanus sp.), mountain-ash (Sorbus sp.) and hackberry (Celtis 

sp.). Members of the maple genus seem to be the preferred host 

of Asian long-horned beetle in Ontario (Kimoto & Duthie-Holt, 

2006; Hu et al. 2009). 


oviPosition or eGG Pits 

Female Asian long-horned beetles use their mandibles to chew 

holes in the bark of hardwood trees. They deposit their eggs in 

these cavities (Kimoto & Duthie-Holt, 2006), also called ovipo- 

Figure 226: Asian long-horned 

beetle with yellow wing spots 50 .

5.2.2 ASIAN LONG-HORNED BEETLE (Anoplophora glabripennis) 

sition pits, which have characteristic 

mandible marks 

(Fig. 227) and range in size 

from 1 to 15mm (Ric et al. 

2007). 

exit holes 

Adult beetles create circular 

holes as they emerge 

after larval development 

is complete (Fig. 228). Exit 

holes ranging in size from 

6 to 12mm can be found 

anywhere on the tree trunk, 

branches and exposed roots 

(Kimoto & Duthie-Holt, 

2006). 

FeedinG GAlleries 

& BArk dAMAGe 

Larval feeding can cause 

separation between the bark 

and sapwood. As a result, 

the bark may appear raised, 

hollow or cracked (Fig. 229). 

After one year, the bark may 

begin to fall off the tree, 

leaving large exposed areas 

with the tunnels or feeding 

galleries visible (Ric et al. 

2007). 

Figure 227: Asian long-horned beetle oviposition pit 

with arrows pointing to mandible marks 51 . 

Figure 228: An exit hole created by an Asian longhorned 

beetle 52 . 

Figure 229: Bark damage 40 (left) and exposed feeding galleries 53 (right). 

[ 193 ]

[ 194 ] 


FrAss or shAvinGs 

Tunneling and feeding larvae leave behind 

a mixture of sawdust and feces that is 

collectively called frass (Fig. 230). Frass 

can sometimes be seen protruding from 

exit holes or cracks in the bark and can 

accumulate at the base of trees or in the 

forks of branches (Kimoto & Duthie-Holt, 

2006). 

Adult FeedinG 

Asian long-horned beetles spend two weeks 

feeding and mating after they emerge from 

trees as adults. They feed on the outer 

tissues of branches, twigs and petioles 

(Fig. 231), and consume the primary and 

secondary veins of leaves (Ric et al. 2007). 

BrAnch dieBAck & hArdwood MortAlity 

High population densities of Asian long-horned beetle can 

weaken and eventually kill hardwood trees. Branch dieback 

caused as a result of larval tunneling occurs from the top 

branches downward, eventually killing the tree (Fig. 232). 

Repeated infestations at lower population densities can produce 

similar results (Nowak et al. 2001). 

Figure 230: Frass created by the 

Asian long-horned beetle 40 . 

Figure 231: Damage caused by adult Asian long-horned beetles feeding on branches 54 

(left) and leaves 40 (right).


Figure 232: Branch dieback 40 (left) and hardwood mortality 40 (right) resulting from Asian long-horned beetle 

infestations. 


syMPtoMs 

whitespotted sawyer (Monochamus scutellatus) 

The whitespotted sawyer is native to North America. Females 

lay their eggs on conifers as opposed to the hardwood species 

that host the Asian long-horned beetle. The wing covers are 

black and appear rough in contrast to the blue-black glossy 

wing covers of the Asian long-horned beetle (Fig. 233). Female 

whitespotted sawyer beetles have white markings on their wing 

covers similar to those of Asian long-horned beetles whereas 

the males are completely black. The easiest way to differentiate 

between the two species is to look for a white or yellow spot 

between the top of the wing covers. Asian long-horned beetles 

do not have that distinctive spot (OMNR, 2010c). 

Figure 233: Comparing the whitespotted sawyer 55 (left) and the Asian 

long-horned beetle 56 (right). 

[ 195 ]

[ 196 ] 


oviposition or egg pits 

Other insects, wildlife and people can cause damage to the outer 

bark of hardwood species that may appear similar to the pits 

created by Asian long-horned beetle. Look for mandible marks 

to determine if an Asian long-horned beetle caused the damage. 

These marks appear on the outer edges of the oviposition pit, 

giving the wound a jagged appearance. Oviposition pits are 

very small, ranging in size from 1 to 15mm (Ric et al. 2007). 

exit holes 

Several species create exit holes similar to those made by adult 

Asian long-horned beetles. Remember that Asian long-horned 

beetles only infest certain hardwood trees, not conifers. The 

gallmaking maple borer (Xylotrechus aceris) and the maple callus 

borer (Synanthedon acerni) are two examples of species that 

create exit holes similar to those of Asian long-horned beetle 

(Fig. 234). Pay attention to the size of the holes. Gallmaking 

maple and maple callus borers create exit holes that are less 

than 5mm in diameter whereas those made by Asian longhorned 

beetles are 6 to 14mm in diameter (Ric et al. 2007). 

Figure 234: Comparing the exit holes of the gallmaking maple borer 57 

(left), to those of the maple callus borer 57 (centre) and the Asian longhorned 

beetle 51 (right). 

Feeding Galleries & Bark damage 

There is an abundance of other insect species that create 

tunnels similar to those made by the Asian long-horned beetle 

in hardwoods. It can be extremely difficult to identify the 

causative agent of feeding galleries just based on tunneling 

patterns. Note the similarities between the tunnels and larvae

(A) 

(B) 

(C) 


of Asian long-horned beetle and those of poplar borer (Saperda 

calcarata), carpenterworm (Prionoxystus robiniae), and red oak 

borer (Enaphalodes rufulus) in Figure 235. Always look for other 

signs and symptoms that can help identify the correct species. 

For further information, see the training guide to detecting the 

signs and symptoms of Asian-long-horned beetle injury written 

by Ric et al (2007). 

Figure 235: (A) Comparing feeding tunnels made by poplar borer larvae 57 

(left) with those made by the Asian long-horned beetle larvae 52 (right); 

(B) comparing the feeding galleries created by the carpenterworm 57 (left) 

with those made by the Asian long-horned beetle 40 (right); (C) comparing 

the feeding galleries created by the red oak borer 58 (left) with those made 

by the Asian long-horned beetle 51 (right). 

[ 197 ]

[ 198 ] 


Frass or shavings 

Frass or shavings are created by many insect borers. Frass can 

be found at the base of hardwood trees or protruding through 

holes and cracks in the bark (Kimoto & Duthie-Holt, 2006). 

Figure 236 shows how frass created by tunneling Asian longhorned 

beetle larvae can easily be confused with that created 

by other insects such as the white oak borer (Goes tigrinus). 

Figure 236: Comparing frass of the white oak borer 57 (left) to that of the 

Asian long-horned beetle 51 (right). 


BioloGy 



The Asian long-horned beetle is 

native to Asia. The first docu- 


mented infestations in North 

....Class. .Insecta 

America occurred in New York 

(1996), Chicago (1998), New Jersey 


(2002 & 2004) and Ontario (2003). 

........Family. Cerambycidae 

Introduction was likely a result of 

having individuals at various life 

...........Genus. Anoplophora 

stages in solid wood packaging 

............Species.. Anoplophora glabripennis. 

material used in international trade 

(Fig. 237). In Ontario, infestations 

were located between the cities of 

Toronto and Vaughan. The species was detected rapidly upon 

introduction and eradication efforts were immediately put into 

place. Approximately 17 000 hardwood trees were destroyed in 

the first year following detection and, although more infested 

trees have since been found and destroyed, the invasion appears 

to be under control (OMNR, 2010c).


Figure 237: Solid wood packing material is a vector for Asian longhorned 

beetle introduction 60 . 

liFe cycle 

The Asian long-horned beetle has a four-stage life cycle including 

egg, larva, pupa and adult (Fig. 238). In Ontario they require 2 

to 3 years to complete their life cycle (Ric et al. 2007). Adult 

females deposit their eggs under the bark of hardwood trees. 

Each female can lay up to 80 eggs during an average adult life 

span of approximately 40 days (OMNR, 2010c). After hatching, 

larvae feed on the inner sapwood and may take 1 to 2 years to 

fully mature. They then enter the pupal stage, usually in June 

and July, and adult beetles emerge approximately 20 days later 

(Ric et al. 2007). 

Egg Larva 


Figure 238: Typical life cycle of the Asian long horned beetle: egg 60 , 

larva 52 , pupa 60 , and adult 49 . 

[ 199 ]

[ 200] 


iMPActs 

The Asian long-horned beetle attacks healthy trees. The larvae 

feed on the inner sapwood creating a series of tunnels that 

effectively block the movement of sap. This can kill the tree 

depending on insect population densities and frequency of 

establishment. The tunnels created in the wood greatly decrease 

timber value (Cartwell, 2007)(Fig. 239). The insect’s preference 

for maple trees is a serious threat to maple syrup operations. 

In addition, large numbers of dead or decaying hardwood trees 

affect the aesthetic value and biological diversity of a forest 

(Dodds & Orwig, 2011). 

Figure 239: Galleries made by Asian long-horned beetle reduce timber 

value 51 . 

Until recently, all Asian long-horned beetle infestations 

occurred in urban areas. In 2008 a new outbreak was reported 

in Worcester, Massachusetts. This outbreak was particularly 

concerning because the infestation was encroaching into a 

hardwood forest. Dodds & Orwig (2011) used this outbreak to 

determine that Asian long-horned beetle can indeed disperse 

rapidly. Their findings have caused a great deal of concern. 

Asian long-horned beetle has the capability of permanently 

altering economically and ecologically important hardwood 

forests throughout North America. 


The Asian long-horned beetle is problematic in parts of its 

native range, particularly in northern China. Populations 

exploded after a large number of suitable host trees, mainly 

in the Populus genus, were planted during a reforestation


campaign on old farmlands. It is hypothesized that such rise in 

abundance greatly enhanced the probability of introductions 

into other countries (Hajek, 2007). 

It is the general consensus that the Asian long-horned beetle 

arrived in North America in solid wood packing material. 

International trade is a common pathway of exotic species 

transportation to new countries. Regulations and inspections 

are mandatory. However, eggs, larvae and even adult beetles 

can be extremely difficult to detect and it is impossible to 

inspect every single product crossing the border (Smith et al. 

2009). Adult beetles can disperse approximately 1 to 3km over 

their lifespan, contributing to the natural spread of established 

populations (Bancroft & Smith, 2005). 



The CFIA established a quarantine zone to control the 

population of Asian long-horned beetles found in the Greater 

Toronto area in 2003. Firewood, live trees, lumber and wood 

chips are prohibited from moving out of the regulated area 

(CFIA, 2012b). Woodlot owners can help prevent the spread of 

Asian long-horned beetles by increasing tree species diversity. 

This has helped suppress outbreaks in China (Hu et al. 2009). 

It is important to avoid introducing firewood and other wood 

products that may harbour eggs, larvae or adult beetles (Smith 

et al. 2009). 


Early detection of Asian long-horned beetle infestations 

requires the regular monitoring of the woodlot by undertaking 

visual inspections of the bark and branches of random 

hardwood trees. Look for various signs and symptoms to avoid 

misidentifications (Turgeon et al. 2010). 


The primary method of control and possible eradication of Asian 

long-horned beetle is to destroy infested trees and all potentially 

infested trees within a given radius (Fig. 240). Removal of all 

potential host trees within a 400m radius around the infested 

tree appears to have successfully controlled the outbreak in 

the Toronto area. The trees were cut and the wood chipped and 

[ 201]

[ 202] 


burned to ensure that all eggs and larvae were killed (Cartwell, 

2007). Approximately 28 700 trees have been removed to date 

from the infested area. The CFIA has offered compensation to 

property owners to replace any trees that were lost as a result 

of the eradication campaign (CFIA, 2012b). 

Figure 240: Removing a tree infested with Asian long-horned beetle 61 . 

Since Asian long-horned beetle is established in North America 

the risk of re-introduction to a quarantined area is high. Thus, 

increased awareness is needed to help detect new infestations 

early. The earlier an infestation is detected the easier it is to 

eradicate and control. Woodlot owners should become familiar 

with the signs and symptoms of an Asian long-horned beetle 

infestation. Report any signs of Asian long-horned beetle to the 

CFIA and/or OMNR as soon as possible (OMNR, 2010c).


5.2.3 

chestnut Blight 

(Cryphonectria parasitica) 


chestnut canker, chestnut bark disease 


Figure 241: Chestnut blight 41 . 

203 

[ ]

[ 204] 


cAusAl orGAnisM 


The fungal pathogen Cryphonectria parasitica is the causal 

agent of chestnut blight disease. Species belonging to the 

genus Cryphonectria are characterized by stromata (i.e., mass 

of fungal filaments and host plant tissues) that are partially 

submerged under the bark of the host tree (Kazmierczak 

et al. 2005). Orange asexual spores, called conidia, have a 

convex shape giving them a swollen appearance. Sexual 

spores, or ascospores, containing partitions created by a single 

septum also characterize species belonging to Cryphonectria 

(Gryzenhout et al. 2006). Cryphonectria parasitica produces an 

abundance of orange conidia and yellow ascospores that may 

at times be seen on the outer bark of the host tree (Fig. 242). 

Underneath the bark, the fungus has a fan-like growth form 

(Horst, 2008). 

Figure 242: Orange fruiting bodies 62 (left) and yellow spores 41 (right) of 

the chestnut blight fungus. 

host(s) 

The two most susceptible host species to the chestnut blight 

fungus are the American (Castanea dentata) (Fig. 243) and 

European chestnut (C. sativa). This exotic disease has hit the 

American chestnut the hardest, whereas in its native range 

has only had a minor effect on four species of chestnut. These 

include Chinese chestnut (C. mollissima), Henry chinkapin 

(C. henryi), Japanese chestnut (C. crenata) and Seguin chestnut 

(C. seguinii), which have all developed a level of resistance to

the disease. There is evidence that the fungus can also grow on 

other hosts such as oak, maple, sumac and hickory. However, it 

causes little to no damage to these hosts (Diller, 1965). 

Figure 243: American chestnut leaves 62 (left) and fruit 5 (right) 

cAnkers 

5.2.3 CHESTNUT BLIGHT (Cryphonectria parasitica) 

siGns And syMPtoMs 

The infection caused by the chestnut blight fungus promotes 

the formation of callus tissue and cankers (Fig. 244). These 

cankers cause the bark to split, creating cracks that can vary 

in size from several centimeters to upwards of a metre (Agrios, 

2005). Cankers may appear orange or yellow due to the presence 

of fungal fruiting bodies and spores (CFS, 2011). 

Figure 244: Stem cankers on young 63 (left) and old 40 (right) 

chestnut trees. 

[ 205]

[ 206] 


wiltinG FoliAGe 

Wilting foliage is a sign of chestnut blight infection (Fig. 245). 

The chestnut blight fungus starts by infecting the bark and outer 

sapwood but eventually spreads over the entire circumference 

of branches and trunks, compromising the vascular system of 

the tree. Water and nutrients can no longer move from the roots 

to the leaves, which results in yellowing and wilting foliage 

(Anagnostakis, 1982). 

Figure 245: Wilting foliage on an American chestnut infected with 

chestnut blight 41 . 

ePicorMic shoots 

Epicormic shoots are often a sign of stress in deciduous 

species. As the chestnut blight fungus spreads around the 

circumference of the trunk it affects the vascular cambium, 

preventing water from reaching the upper branches and leaves. 

In response, the tree sends out epicormic shoots underneath 

the cankers and dead tissue (Fig. 246) (Anagnostakis, 1987). 

Figure 246: Epicormic shoots growing below a chestnut blight-induced 

canker 41 .


syMPtoMs 

There are several species of parasitic fungi belonging to the 

genus Nectria that can cause similar symptoms to those of 

chestnut blight. These species of Nectria infect hardwood trees 

and create target-like cankers or large areas of callus tissue (Fig. 

247). Some Nectria species may infect chestnut trees but these 

cankers are rarely fatal (Brandt, 1964). Cankers and cracks may 

also form on the trunks and branches of American chestnut 

as a result of frost damage or sunscald. This damage usually 

occurs as a result of fluctuating temperatures (i.e., as the water 

freezes and thaws, exerting pressure on the outer bark) and 

callus tissue is produced by the tree in response to the damage 

(Swift et al. 2008). 


Kingdom. Fungi 

..Division. Ascomycota 

....Class. Sordariomycetes 

......Order. Diaporthales 

........Family. Valsaceae 

..........Genus. Cryphonectria 

............Species. Cryphonectria parasitica 


Figure 247: Target-like cankers 41 (left) and excessive callus tissue (right) 41 on native hardwoods resulting 

from Nectria infection. 

BioloGy 


The chestnut blight fungus was 

likely introduced into North America 

on nursery stock from Asia. It was 

first discovered in New York in 

1904 when American chestnut trees 

started deteriorating (Diller, 1965). 

The disease spread very quickly and 

within a period of 50 years it was 

present across the entire range of the 

American chestnut. It is estimated 

that 3.5 billion American chestnut 

[ 207]

[ 208] 


trees have been killed by this pathogen (Turchetti & Maresi, 

2008). The first reports of the disease in Ontario occurred in 

the 1920’s. Today, less than 700 trees have been documented 

in the Carolinian zone of southern Ontario, the American 

chestnut’s natural range (Fig. 248). The majority of surviving 

trees occur in the forest understory as non-reproductive 

individuals (Tindall et al, 2004). Chestnut blight has also been 

introduced to Europe where it is causing similar devastation on 

the European chestnut. However, evidence of natural recovery 

from blight symptoms has been observed since the early 1950’s 

(Brewer, 1995). For more information on natural recovery see 

discussion on hypovirulence in control options. 

Figure 248: Natural range of the American chestnut (C. dentata) 64 . 

diseAse cycle 

The fungal pathogen C. parasitica overwinters within the 

sapwood of its host tree. In the spring, ascospores (sexual 

spores) and conidia (asexual spores) are released into the 

environment (Tattar, 1978). Ascospores are released and 

transported to new hosts via wind currents while conidia are 

transported with the help of rainwater, animals and insects 

(Kazmierczak et al. 2005). Spores are dispersed throughout the 

year, with wet weather enhancing dispersal ability (Sinclair & 

Lyon, 2005). Spores infect new hosts through wounds in the 

bark and, once there, germinate and develop filaments (i.e., 

hyphae) that invade the healthy tissue of the inner bark and 

cambium. A structure called a stroma is formed that houses 

the fruiting bodies in which spores are formed (Kazmierczak et 

al. 2005). Figure 249 demonstrates the disease cycle of chestnut 

blight.


Formation.of.fruiting.bodies.. 

and.spores 41 . 

Cankers.are.formed.at.sites.. 

of.infection 63 .. 

.............Spores.transported.by.wind,.rain.or. 

.................other.organisms.to.new.hosts 66 . 

..............Spores.infect.host.trees. 

...........through.wounds.in.the.bark 67 . 

Figure 249: Disease cycle of chestnut blight. 

iMPActs 

The greatest impact resulting from the introduction of chestnut 

blight is its devastating effects on the American chestnut. 

Both the provincial and federal governments have listed the 

American chestnut as an endangered species (Environment 

Canada, 2011; OMNR, 2012b). The loss of the vast majority 

of large American chestnuts in southern Ontario has had an 

impact on forest biodiversity. Prior to the introduction of the 

disease, southern Ontario had as many as 2 million American 

chestnut trees. Today less than 700 trees have been documented 

in Ontario (Tindall et al. 2004). Chestnut blight does not kill 

the root system thus allowing the American chestnut to escape 

extinction by existing as small saplings in the forest understory 

(Fig. 250). However, there is little possibility of these saplings 

becoming established because they quickly become infected 

with age (Sinclair & Lyon, 2005). 

Figure 250: American chestnut surviving in the forest understory 27 . 

[ 209]

[ 210] 



Spores of the chestnut blight fungus are constantly being 

released from infected trees. Ascospores (sexual spores) 

are transported long distances via wind currents. Conidia 

(asexual spores) are transported shorter distances with the 

help of rainwater and animal or insect vectors (Manion, 1991). 

The fungus can survive even after the host tree has died. In 

fact, it has been observed growing on fallen logs. As such, the 

movement of firewood could contribute to the spread of this 

devastating disease (Diller, 1965). Other tree species, such as 

oaks, maples, sumacs and hickories, may also act as hosts 

allowing the fungus to remain viable in the forest even after 

the loss of the American chestnut as a primary host. These 

genera have varying degrees of susceptibility to the fungus; 

only the American chestnut appears to die as a result of the 

infection (Tindall et al. 2004). 



Keeping a diverse and healthy woodlot is the best means of 

alleviating the effects of chestnut blight. Strong and healthy 

chestnut trees have a better chance of resisting infection or 

recovering should they become infected (Boland et al. 2000). 

Competition from other forest trees, heavy frosts, and the 

presence of other insects and pathogens increase the damage 

caused by chestnut blight infections (Griffin, 2000). The 

American chestnut is shade intolerant and requires high light 

levels for optimal growth (Fig. 251). Since healthy trees may 

be less susceptible to infection, consider removing any trees 

that are directly competing with American chestnut. This will 

promote vigorous growth and contribute to the overall health 

of the tree (Boland et al. 2000). 

Figure 251: A healthy American chestnut leaf 5 .


It is good practice to limit the movement of firewood that may 

act as a vector for spreading the disease. Be especially careful 

during woodlot management activities not to damage or wound 

trees as chestnut blight spreads through airborne or organismtransported 

spores that can infect healthy trees through 

wounds in the bark (Boland et al. 2000). 


It is important to know if American chestnut is present in your 

woodlot. Woodlot surveys are encouraged to identify the species 

composition of the forest. If American chestnut is present, it is 

recommended that these individuals are monitored for signs of 

chestnut blight infection. Becoming familiar with the American 

chestnut trees in your woodlot will help you to recognize any 

new signs or symptoms of infection (Boland et al. 2000). Pay 

particular attention to the small branches. Infection usually 

occurs in smaller branches before it spreads to larger, more 

prominent ones. Wilting foliage and branch dieback is another 

symptom to be on the lookout for (Sinclair & Lyon, 2005). 


There are several ways in which the chestnut blight 

epidemic is currently being handled in Ontario, including 

the implementation of forest management practices, the 

incorporation of hypovirulence, and the development of blightresistant 

genotypes. 

American chestnut has not been completely eliminated from 

North America’s forests because the rootstocks are not affected 

by the disease. Once infected, the aboveground portion of 

the tree will die off leaving an uninfected rootstock that will 

periodically send up new shoots. However, these emerging 

shoots are not immune to the disease and usually become 

blighted with age. Rootstocks may eventually succumb to the 

continuous pressure of diseased stems (Turchetti & Maresi, 

2008). Thus, it is important to keep these understory saplings 

healthy by alleviating environmental stresses and promoting 

blight resistance through proper forest management (Sisco, 

2012). 

American chestnut is a shade intolerant species that benefits 

from thinning practices. Harvesting a small portion of the 

trees in the woodlot to create openings in the canopy and 

discourage competition among desirable species will allow 

the American chestnut to establish. Cultural practices such 

as removing infected twigs and branches are recommended 

to lower the fungal population in the area. These infected 

branches should be promptly bagged and incinerated (Boland 

et al. 2000). Remember that American chestnut is considered 

[ 211]

[ 212] 


an endangered species protected by the Endangered Species 

Act, 2007. Consult a species-at-risk biologist from the OMNR 

before removing an American chestnut tree. 

In the early 1950’s, European chestnuts in Italy were observed 

to be healing from the chestnut blight symptoms (Brewer, 1995). 

The chestnut blight fungus contained a virus that made the 

blight pathogen less virulent to their hosts. This type of natural 

recovery has been termed hypovirulence (Barakat et al. 2009). 

American chestnut trees with hypovirulence exhibit superficial 

cankers that affect the outer bark while leaving the vascular 

cambium intact. This ultimately allows them to survive an 

otherwise deadly disease (Brewer, 1995). Hypovirulence has 

since been documented to occur in American chestnuts in 

several locations in North America (Griffin et al, 1983; Brewer, 

1995) including Ontario, Canada (Melzer & Boland, 1999; Tindal 

et al. 2004). Researchers have been attempting to introduce 

the hypovirulent strains throughout the American chestnut’s 

range, however with limited success. Natural dispersal of the 

hypovirulent strains is almost non-existent. This may be due 

to reduced spore production as well as the inability for certain 

virulent strains to fuse successfully with the hypovirulent 

strains (Manion, 1991; Griffin, 2000). 

Research on genetic engineering and the creation of transgenic 

trees (i.e., American chestnut containing blight-resistant genes 

from Chinese chestnut) is currently underway. These potentially 

blight-resistant genes may help the American chestnut gain 

resistance. However, introducing transgenic trees to the forest 

ecosystem is a matter of debate (Hirsch, 2012). 

Another method currently being investigated is interspecific 

hybridization. Creating blight-resistant Chinese/American 

chestnut hybrids using a method of backcross breeding is 

currently underway in Canada by the Canadian Chestnut 

Council (www.canadianchestnutcouncil.org) and in the 

United States by the American Chestnut Foundation (www.acf. 

org). The goal of backcross breeding is to create blight-resistant 

trees that are morphologically similar to the American chestnut. 

Hybrids of Chinese and American chestnut are bred over three 

generations with pure American chestnuts. This breeding 

method creates trees that will appear in every aspect similar to 

American chestnut with the added benefit of blight resistance 

(Diskin et al. 2006). It is expected that the small percentage of 

exotic genes will provide the same level of natural resistance 

as that observed in chestnut species that co-evolved with C. 

parasitica (Griffin, 2000). Field trials are currently underway to 

test the level of blight-resistance in natural forest ecosystems 

(Smith, 2012).


5.2.4 

Beech Bark disease complex 

(Cryptococcus fagisuga & Neonectria spp.) 


BBd, Beech bark canker, neonectria canker 


Figure 252: Beach bark disease 62 . 

213 

[ ]

[ 214 ] 


cAusAl orGAnisMs 


Beech bark disease is caused by the combined effects of an insect 

and a pathogen. The beech scale insect (Cryptococcus fagisuga) 

is very small, only reaching lengths of approximately 1mm (Fig. 

253). These tiny insects feed and lay their eggs on American 

beech (Fagus grandifolia) and European beech (F. sylvatica). The 

larvae form a protective white, waxy coating that hides them 

from view (Fig. 254). In heavy infestations this substance can 

cover large portions of the trunk (Wainhouse, 1980). Feeding 

by the scale insect kills cells in the outer bark tissues creating 

wounds in which pathogen infection can occur (Ehrlich, 1934). 

Figure 253: Beech scale (Cryptococcus fagisuga) 41 . 

The primary fungal pathogen 

associated with beech bark 

disease is the invasive 

Neonectria faginata. A native 

fungus, N. ditissima, can 

also infect beech after scale 

colonization, however severe 

infections are rare in Ontario 

(McLaughlin & Greifenhagen, 

2012). Both species form 

red fruiting bodies, called 

perithecia, which can be seen 

on the outer bark (Fig. 255). 

Spores are released from the 

perithecia and transported to 

new hosts via wind and rain 

(Houston & O’Brien, 1983). 

Figure 254: White larval secretions created by beech 

scale 68 . 

Figure 255: Fruiting bodies of Neonectria species 69 .

5.2.4 BEECH BARK DISEASE (Cryptococcus fagisuga & Neonectria spp.) 

host(s) 

Two species of beech are primarily affected by beech bark 

disease in Ontario: American beech (Fagus grandifolia), which 

is native to North America and European beech (F. sylvatica). 

American beech is shade tolerant and typically occurs in climax 

forests. Conversely, European beech is commonly planted as 

an ornamental in Ontario. The leaves of American beech have 

9-14 vein pairs whereas European beech has 5-9 vein pairs (Fig. 

256). The teeth on the margins of American beech are more 

prominent than those of European beech (Farrar, 1995). 

Figure 256: Comparing the leaves of American beech 1 (left) to those of European beech 15 (right). 

white, wAxy wool 


The first sign of beech bark disease is the occurrence of beech 

scale, whose larvae secrete a white wool-like substance (Fig. 

257). As insect populations increase the tree becomes covered 

with white larval secretions and in large infestations the main 

stem can appear almost entirely white (Wainhouse, 1980) (Fig. 

258). 

Figure 257: Scale insect producing a white, waxy 

wool 31 . 

Figure 258: American beech covered in larval 

secretions from beech scale 41 . 

[ 215 ]

[ 216] 


necrotic sPots 

Infection by Neonectria fungi causes necrosis of the bark and 

inner sapwood, which may produce a reddish-brown sap (Fig. 

259). As the bark continues to decay and the infection spreads, 

beech scale populations decrease. The fungus eventually 

destroys the bark, cambium and outer sapwood, leaving trees 

vulnerable to other insects and diseases (Sinclair & Lyon, 2005). 

Figure 259: Necrotic spots on American beech 41 . 

cAnkers & cAllus tissue 

Cankers form on the main stem and branches as a result of 

beech bark disease (Fig. 260). A natural defense of the tree is 

to create callus tissue. However, after 2 to 3 years of infection, 

large pieces of bark begin to fall off, revealing areas of extensive 

decay (Sinclair & Lyon, 2005) 

Figure 260: Cankers and callous tissue resulting from beech bark disease 71 .



syMPtoMs 

Phytophthora bleeding cankers 

Several species of fungal pathogens in the Phytophthora genus 

can create cankers similar to those caused in response to 

beech bark disease. These cankers produce a reddish-brown 

substance giving the appearance that the trunk is bleeding. 

These cankers may at times appear similar to the necrotic 

spots caused by beech bark disease (Fig. 261). Look for the 

white, wooly signs of the beech scale to help determine whether 

the necrotic spots are indeed a result of beech bark disease. 

Some Phytophthora bleeding cankers will cause tree mortality. 

Consult a professional arborist or forester should these types 

of canker appear (Pijut, 2006). 

Figure 261: Comparing Phytophthora bleeding cankers 33 (top) and 

necrotic spots resulting from beech bark disease 41 (bottom). 

[ 217 ]

[ 218] 



Beech.Scale.Insect. 




......Order. Hemiptera 

........Family. Eriococcidae 

..........Genus. Cryptococcus 

............Species. Cryptococcus fagisuga. 

Neonectria fungi. 

BioloGy 


..Phylum. Ascomycota 


......Order. Hypercreales 

........Family. Nectriaceae 

..........Genus. Neonectria 

............Species. Neonectria ditissima. 





......Order. Hypercreales 

........Family. Nectriaceae 

..........Genus. Neonectria 

............Species. Neonectria faginata 

Beech scale was introduced to North America in the 1890’s. It 

was first observed in Nova Scotia and is thought to have arrived 

on ornamental stock of European beech (Houston & O’Brien, 

1983). In 1981, the first evidence of the beech scale in Ontario 

was found in Newmarket as a result of an investigation into the 

decline of beech trees in the area (Bisessar et al. 1985). However, 

it was not until 1999 that the beech bark disease complex was 

officially confirmed in Ontario. Today, beech bark disease is 

present across the majority of the native range of American 

beech (McLaughlin & Greifenhagen, 2012) (Fig. 262) 

diseAse cycle 

In the summer months, adult scale insects lay their eggs on 

the outer bark of beech trees. Upon hatching the nymphs can 

disperse to new hosts via wind and animal vectors (McLaughlin 

& Greifenhagen, 2012). In the late summer and fall, the nymphs 

attach to the outer bark and start producing a protective woolly 

coating. These immobile nymphs hibernate over the winter and 

moult into adults in the spring (Shigo, 1972). While feeding, the 

beech scale insects create wounds in the host tissue, increasing 

the tree’s susceptibility to pathogen infection. Fungal spores of 

Neonectria species are carried by wind or water from infected 

trees to new hosts where they can easily establish and germinate 

within tiny wounds in the bark (McLaughlin & Greifenhagen, 

2012).


Figure 262: Natural range of American beech (Fagus grandifolia) 64 . 

Beech bark disease has three stages (Fig. 263). The first stage is 

called the advancing front. This is when scale insects become 

established in a forest stand and begin to reproduce on the outer 

bark of beech trees. The next stage or killing front is when the 

Neonectria species infect the tree, resulting in high levels of 

mortality. The aftermath forest consists of a few scattered large 

beech trees and many infected young saplings (Shigo, 1972). 

Advancing front:.Invasion.by. 

beech.scale 68. (top). 

Killing front:.Disease.complex. 

causes.tree.mortality 41 (top.right). 

Aftermath forest:.Few. 

surviving.large.beech.trees, 

forest.characterized.by.small 

saplings 41. (bottom.right).. 

Figure 263: Stages of beech bark disease. 

[ 219 ]

[ 220] 


iMPActs 

American beech is an important food source for wildlife. It 

produces a large nut crop every 2 to 8 years and provides habitat 

for birds and mammals (Tubbs & Houston, 1990). By causing 

high levels of beech mortality beech bark disease results in the 

sudden loss of an abundant and important resource in forest 

food webs (Storer et al. 2005). Beech bark disease may also alter 

the species composition of the forest through indirect effects 

that inhibit the regeneration of native hardwood species. As 

large beech trees succumb to the disease they send up multiple 

saplings that can quickly form dense patches of beech saplings 

that compete with other regenerating species (Nyland et al. 

2006). 

American beech wood is used to make flooring and furniture. 

Therefore, compared to others, woodlots with a high percentage 

of beech trees may be at an increased risk of economical losses as 

a result of beech bark disease (Farrar, 1995). Beech bark disease 

often girdles the tree and causes structural weakening up to 

the point of collapse (Fig. 264). The tops of large beech trees 

have been observed to break off the main stem, an occurrence 

known as “beech snap”. It can be a hazard to humans, especially 

in recreational areas (Heyd, 2005). 

Figure 264: Tree mortality caused by beech bark disease 41 . 


The beech scale insect is the first stage of the disease complex. 

These tiny insects can disperse during what is called a crawler 

stage. At this stage the insects have some mobility and are 

largely aided by wind currents. Scale insects have been observed


to disperse up to 10m to a new host tree (Wainhouse, 1980). 

Humans are also thought to play a major role in the dispersal 

of scale insects by moving ornamental stock and firewood 

(McLaughlin & Greifenhagen, 2012). Neonectria spores are 

vectored through wind currents and water to new host trees 

where infection occurs at wound sites created by the feeding of 

scale insects (Shigo, 1972). 



Consider implementing preventative silvicultural techniques 

in uninfested woodlots by increasing tree species diversity. 

This can be accomplished through selective thinning practices 

(McCullough et al. 2005). American beech is a desirable species 

that contributes to diversity and is an important source of 

food and habitat for wildlife (Storer et al. 2005). As such, the 

complete removal of beech trees is not recommended. Choose 

to retain large healthy beech trees with smooth bark while 

harvesting those with signs of decay or injury, which are more 

likely to be infected (McCullough et al. 2005). Moving firewood 

increases the risk of introducing beech bark disease into new 

areas (Jacobi et al. 2011), especially from mid-summer to late 

fall when the scale insects are relatively mobile (Heyd, 2005). 


Early detection requires regular monitoring and surveying the 

woodlot for signs of beech bark disease. The first signs of the 

advancing front are the characteristic white wooly substance 

produced by the beech scale insect (Shigo, 1972). Look for beech 

trees with rough bark as the scale insects seem to prefer the 

rougher texture, which may provide greater protection from 

enemies or the weather. Look at larger older trees; these seem 

to be the first to be colonized by scale insects (Koch & Carey, 

2005). If the beech scale is present, the next sign to look for is 

the formation of necrotic spots and cankers. As the Neonectria 

fungus begins to destroy the tree scale insects die and the 

white wooly cover turns black. This is a sign that the killing 

front is advancing (Shigo, 1972). 


Prevention strategies are available to help protect a forest 

from beech bark disease. However, there is no absolute way of 

ensuring that a woodlot will remain disease free. When faced 

[ 221]

[ 222] 


with the advancing front of beech bark disease there are several 

ways in which to alleviate damage. Conduct a survey of the 

woodlot to identify the extent of beech scale infestation. High 

value trees may be washed off with water in an attempt to save 

the tree from fungal infection. However, this is only practical for 

a small number of trees as regular washings will be needed to 

prevent re-colonization (McCullough et al. 2005). 

Identify potentially resistant trees that either have a very small 

amount of beech scale or none. These trees’ genotypes should 

be protected since resistance is extremely rare. Approximately 

1% of all the beech trees in North America are thought to be 

resistant to beech bark disease, thus it is important to keep 

resistant genes in the gene pool (McLaughlin & Greifenhagen, 

2012). Research is underway to increase the number of diseaseresistant 

trees in natural areas. However, breeding for resistance 

has proven difficult because beech does not reach a reproductive 

age for approximately 40 years, at which time they are nearly 120 

feet tall. Despite these challenges, recent studies have shown 

that disease resistance is heritable (Koch & Carey, 2004) and that 

a high number of seedlings derived from resistant parents will 

also show signs of resistance (Koch et al. 2010). 

Consider harvesting or thinning 

the woodlot to salvage 

heavily infected trees. Diseased 

saplings may undergo 

vigorous growth in response 

to higher light levels resulting 

from thinning practices. 

However, these saplings may 

compartmentalize cankers 

and grow with trunk defects 

that greatly decrease their 

timber value (Fig. 265). Removing 

these deformed saplings 

will provide room for other 

species to grow and contribute 

to the overall diversity of 

the woodlot (McCullough et al. 

2005). Beech also has the ability 

to multiply via vegetative 

reproduction. Numerous saplings 

may emerge from underground 

roots associated with 

infected trees. These saplings 

should be removed as they 

will eventually become infected 

and create competition 

for other understory species 

(Smallidge & Nyland, 2009). 

Figure 265: American beech growing 

with deformities 66 .


5.2.5 

Gypsy Moth 



european gypsy moth, Asian gypsy moth 


Figure 266: Gypsy moth larva 72 . 

223 

[ ]

[ 224] 


orGAnisM 


Gypsy moths range from 2.5 to 3cm in length. Males have light 

brown wings with darker coloured markings. Females are much 

lighter in colour. They have creamy white wings with markings 

that can be either distinctive or faint (Fig. 267). Males have large 

grayish-brown feathery antennae (Benoit & Lachance, 1990) 

(Fig. 268). There are two exotic strains of gypsy moth present 

in North America, one from Europe and one from Asia. Only the 

European strain has been documented in Ontario. All females 

of the European strain are flightless (Nealis & Erb, 1993). 

Figure 267: Comparing female (left) and male (right) 

gypsy moths 34 . 

Figure 268: Male gypsy moth with characteristic 

feather-like antennae 68 . 

Gypsy moth caterpillars go through several developmental 

stages (Fig. 269). During the 1 st stage, called 1 st instar, the 

caterpillar appears reddish-brown and is covered in fine black 

hairs. During the 2 nd and 3 rd instars the body turns to a mottled 

black and yellow colour. Orange and gray spots appear along 

the back. The later instars have a central stripe showcasing 

5 rows of blue spots and 6 rows of red spots. Prominent tufts 

of hair cover the body (McManus et al. 1989). 

Figure 269: Changes in appearance from early 74 (left) to later 75 (right) instar development of 

gypsy moth larvae.

host trees 

The European strain of gypsy moth present in Ontario has a 

wide host-range, which includes over 300 species of hardwood 

trees. However, the most commonly preferred trees are oak 

(Quercus sp.), willow (Salix sp.), aspen (Populus sp.) and birch 

(Betula sp.). Only the larvae cause damage to these host trees 

through feeding on the leaves (Tobin & Liebold, 2011). The 

Asian strain has an even wider host range than the European 

strain, feeding on both hard and softwood species (Régnière et 

al. 2009). 

eGG MAsses 


Female gypsy moths create large, light brown egg masses, 

which can be found in sheltered areas such as sheds, under 

picnic tables, on vehicles, etc. (McManus et al. 1989) (Fig. 270). 

During outbreaks, multiple egg masses can be seen on the 

tree trunks and branches of hardwood trees. Each egg mass 

is approximately 3.5 x 2cm and can contain up to 1000 eggs 

(Nealis & Erb, 1993). 

lArvAl FeedinG 

5.2.5 GYPSY MOTH (Lymantria dispar) 

Figure 270: Egg masses of gypsy moth are found both outdoors 76 (left) and in sheltered locations 77 (right). 

Gypsy moth larvae, or caterpillars, in Ontario feed on the leaves 

of various hardwood species. They avoid the veins and midveins, 

creating leaves with a skeletal appearance (Fig. 271). 

However, during severe outbreaks, complete defoliation can 

occur. Larvae feed mostly at night and hide in communal areas 

during the day (Nealis & Erb, 1993). 

[ 225]

[ 226] 


Figure 271: Gypsy moth larvae feeding on leaves 78 (left) and congregating on a tree trunk 43 (right). 


syMPtoMs 

eastern tent caterpillar (Malacosoma americanum) 

The eastern tent caterpillar is a native defoliating insect that 

has similar population outbreaks to those of the gypsy moth. 

Larvae of both species feed at night and hide during the day. 

However, eastern tent caterpillars construct silken tents in 

which they congregate and find shelter. Eastern tent caterpillars 

have a white stripe bordered in yellow with blue markings along 

their sides (Rabaglia & Twardus, 1990) (Fig. 272). 

Figure 272: Eastern tent caterpillar 39 (left) and a silken tent 40 (right).

Forest tent caterpillar (Malacosoma disstria) 

Forest tent caterpillars are native to North America. Similarly 

to gypsy moth, the larvae feed at night and congregate during 

the day, often in large clumps on tree trunks. The forest tent 

caterpillar has a row of alternating small and large white spots 

along their backs that give the impression of footprints (Fig. 

273). Gypsy moth caterpillars have five blue and six red dots 

along their backs (Dodds & Seybold, 1996). 

Figure 273: Forest tent caterpillar 79 (left) and a large congregation 44 (right). 



BioloGy 



The European strain of gypsy moth was 

introduced into North America in 1869. 


Several moths escaped from the office of 


an entomologist, Leopold Trouvelot, who 

was conducting crossbreeding research 

......Order. Lepidoptera with silk worms in Massachusetts 

........Family. Lymantriidae 

(Benoit & Lachance, 1990). Within 20 

years severe defoliation brought the 

..........Genus. Lymantria 

gypsy moth to public attention (Tobin & 

............Species. Lymantria dispar. 

Liebold, 2011). It is unknown when the 

species established in Canada but the 

first outbreak requiring management 

occurred in Quebec in 1924 (Nealis, 2002). The European strain 

was first observed in Ontario in 1969 whereas the Asian strain 

has become established in western North America and has not 

been detected in Ontario (Nealis & Erb, 1993). 

[ 227]

[ 228] 


liFe cycle 

Gypsy moths have a four-stage life cycle, including egg, larvae, 

pupae, and adult (Fig. 274). In the fall female moths create 

large egg masses that contain up to 1000 eggs. Eggs overwinter 

in these masses and hatch the following spring. Larvae, or 

caterpillars, emerge from their eggs and begin feeding on 

leaves from May through June. After feeding, they enter the 

pupal stage in which the larvae undergo metamorphosis from 

a caterpillar to a moth. This process takes approximately 2 

weeks. Upon emerging from cocoons the adult moths mate and 

each female creates a single egg mass (Nealis & Erb, 1993). 

Figure 274: Life cycle of the gypsy moth including egg 76 , larvae 40 , 

pupae 47 and adult 48 life stages. 

iMPActs 

Egg Larva 


The introduction of gypsy moth to North America had 

devastating effects on deciduous forests. Larvae can completely 

defoliate entire forest stands during an outbreak. Defoliation 

has been linked to both hardwood growth loss and mortality, 

with forest stands experiencing 25-30% mortality as a result 

of an outbreak (Fig. 275). In areas with already stressed trees 

mortality can reach 100% (Tobin & Liebold, 2011). 

Figure 275: Defoliation caused by a gypsy moth outbreak 80 .


Economic hardships ensue with the loss of hardwood trees. 

For instance, defoliation and mortality affect the value of 

timber products. In areas affected by gypsy moth outbreaks, 

quarantines are often implemented to prevent further spread. 

These quarantines have economic impacts associated with 

trade and the movement and sale of wood products. Tourism 

may also be affected as the aesthetic value of natural forested 

areas is compromised (Régnière et al. 2009). 


Flightless adult females characterize the 

European strain of gypsy moth presently 

found in Ontario (Nealis, 2002). They release 

a pheromone to attract the males and do not 

disperse from the tree or structure in which 

they reached the pupal stage (Nealis & Erb, 

1993). Dispersal occurs during the larval 

stage. Young caterpillars climb to the upper- 

most branches of the canopy and hang from 

long silken threads. Winds can then aid in 

their movement to new host trees, a type 

of passive dispersal known as ballooning 

(Hajek & Tobin, 2010). 

Long distance dispersal occurs via human 

vectors (Fig. 276). Females deposit egg 

masses in sheltered locations near the 

ground, often in man-made structures. For Figure 276: Female gypsy moths 

ovipositing on a tire 

instance, egg masses have been found in 

vehicles, campers, tents and lawn furniture (Régnière et al. 

2009), with firewood being one of the leading causes of longdistance 

dispersal (Bigsby et al. 2011). 

81 . 



Preventing movement at all stages of the gypsy moth’s lifecycle 

is key. Currently, outbreaks occur throughout southern 

Ontario, up to and including Sault Ste. Marie. Quarantines are 

enforced by the CFIA to help prevent further spread. Before 

moving to areas outside of the quarantine zone it is important 

to inspect vehicles and outdoor equipment for egg masses, 

larvae, cocoons or adult moths (CFIA, 2011a). 

[ 229]

[ 230] 



Monitoring is the first step to detecting 

new or re-occurring infestations. Inspect 

all outdoor equipment for egg masses. 

Placing a burlap sack around the lower 

trunk of a preferred host tree such as oak 

may attract gypsy moth larvae as it provides 

shelter during the day (Fig. 277). Regularly 

monitoring the contents of the sack can not 

only enable early detection but also inform 

the woodlot owner as to the state and 

severity of infestation. Any trapped gypsy 

moth caterpillars should be destroyed to 

help reduce local populations (McManus et 

al, 1989). 


Eradication programs involving the gypsy moth in North 

America have not been successful. The goal has changed to 

suppression and control. The CFIA enforces quarantines in 

infested areas to prevent further spread. Pheromone traps are 

used as a monitoring tool outside of the quarantine zones. 

Outbreaks within the quarantine zone are the responsibility 

of municipalities and/or private woodlot owners (Régnière et 

al. 2009). 

Woodlot owners are encouraged to actively manage gypsy moth 

infestations on their property. Performing a regular survey of 

outdoor equipment for egg masses is an important first step 

because each can contain up to 1000 eggs. Egg masses should 

be destroyed by placing them in hot, soapy water or by burning. 

Attention: the hairs of gypsy moth caterpillars and fibers 

from egg masses may cause allergic reactions in some people 

(McManus et al. 1989). 

Maintaining the health of the woodlot is another important 

factor when dealing with re-occurring gypsy moth infestations; 

healthy trees are less likely to die as a result of defoliation. 

Good woodlot management practices help to increase habitat 

for wildlife and promote populations of natural gypsy moth 

predators such as small mammals and birds (Nealis & Erb, 1993). 

Pesticides may be used in conjunction with other management 

techniques. The most commonly used pesticide for gypsy moth 

control is Bacillus thuringiensis (Btk), which must be applied by 

a licenced exterminator. This pesticide consists of a bacterium 

that produces protein crystals that are toxic to gypsy moth 

larvae (Hajek & Tobin, 2010). Although it should be taken into 

consideration that Btk is also toxic to native caterpillars, there 

is evidence that negative impacts are confined to the first year 

of Btk application (Solter & Hajek, 2009). 

Figure 277: Using a burlap sack as 

a method to attract gypsy moth 

larvae 82 .


5.2.6 

dutch elm disease 



ded 


Figure 278: Elm affected by Dutch Elm Disease 83 . 

231 

[ ]

[ 232] 


cAusAl orGAnisMs 


Dutch elm disease is caused by three species of ascomycete 

fungi that cut off sap movement in elm trees (Ulmus spp.). 

These species are Ophiostoma ulmi, O. novo-ulmi, and O. himalulmi. 

These pathogenic fungi belong to a group called the 

Ophiostoma piceae complex, which is characterized by fungi 

with fruiting bodies containing two distinct spore-bearing 

structures (Harrington et al. 2001): black perithecia (i.e., fruiting 

body containing spores) and orange asexual spores (Chung et 

al. 2006)(Fig. 279). 

Figure 279: Fruiting bodies and asexual spores of Ophiostoma ulmi 84 . 

hosts 

Dutch elm disease affects all three native species of elm, 

including white elm (Ulmus americana), slippery elm (U. rubra), 

and rock elm (U. thomasii) (Fig. 280). The disease also affects 

two introduced elms: wych elm (U. glabra) and English elm (U. 

procera). Conversely, Siberian elm (U. pumila) is an introduced 

ornamental that has shown resistance to Dutch elm disease 

(Farrar, 1995).

(A) 

(B) 

(C) 

5.2.6 DUTCH ELM DISEASE (Ophiostoma spp.) 

Figure 280: Leaves and native range distribution 64 of white elm 5 (A), slippery elm 5 (B) and rock elm 5 (C). 

[ 233]

[ 234] 


wiltinG leAves 


The first signs of infection usually appear in early summer. 

The leaves on isolated branches turn yellow and begin to wilt 

(Fig. 281). As the infection spreads and more branches wilt, the 

crown will show bare areas where the leaves have wilted and 

fallen off the tree. Eventually the entire crown becomes infected 

and the tree dies. Tree mortality usually occurs between 1 to 3 

years after infection (Davis & Meyer, 1997). 

Figure 281: Wilted foliage as a result of Dutch elm disease 41 . 

sAPwood discolourAtion 

Signs of infection, which usually begins within individual 

branches, can be seen in the sapwood. As the fungus spreads 

the sapwood turns dark brown (Fig. 282). Cutting a cross section 

of the inner wood will reveal a dark brown ring indicating the 

presence of the fungus. Peeling off the bark will also reveal a 

dark brown streaking on the outer sapwood (Sinclair & Lyon, 

2005). 

Figure 282: Streaking shown in a cross section 85 (left) and on the outer sapwood of an elm branch 59 (right).


siMilAr diseAses, siGns 

& syMPtoMs 

elm yellows (Candidatus Phytoplasma spp.) 

Elm yellows disease is caused by phytoplasmas (i.e., small 

specialized obligate parasitic bacteria of plant tissue that lack 

a cell wall) (Lee et al. 2004). As with Dutch elm disease, it also 

affects the vascular system of elm trees causing the leaves to 

turn yellow and wilt (Fig. 283). However, the entire crown wilts 

simultaneously whereas Dutch elm disease starts in individual 

branches and gradually spreads through the entire crown. Elm 

yellows will also cause discolouration of the inner sapwood but 

it produces a distinct wintergreen odour (Sinclair, 2000). 

Figure 283: Comparing the symptoms of elm yellows 40 (left) and Dutch elm disease 65 (right). 

verticillium wilt (Verticillium spp.) 

Soil-borne fungi that affect the vascular system of trees and 

shrubs causes Verticillium wilt. This disease can be confused 

with Dutch elm disease because it causes similar yellowing of 

leaves and sapwood discolouration (Fig. 284). However, even 

though Verticillium wilt may cause tree mortality, it is often 

much less severe than Dutch elm disease. Cankers may also 

form as a response to Verticillium wilt, which is a symptom 

associated with Dutch elm disease (Ash, 1994). 

Figure 284: Leaf wilting 41 (left) and sapwood discolouration 69 (right) 

associated with Verticillium wilt. 

[ 235]

[ 236] 



BioloGy 



Dutch elm disease is caused by three 

species of pathogenic fungi. Ophiostoma 

..Division. Ascomycota 

ulmi was the first of the three species 


to cause widespread mortality. It was 

......Order. Ophiostomatales 

introduced to Europe around 1910 and 

........Family. Ophiostomataceae subsequently to North America in the 

..........Genus. Ophiostoma 

1940’s (Brasier, 1990; 1991). In Ontario, 

the first reports of Dutch elm disease 

Species Ophiostoma ulmi 

date from 1946, two years after having 

Ophiostoma novo-ulmi 

been found in Quebec (Hubbes, 1999). 

Ophiostoma himal-ulmi 

Ophiostoma ulmi has been described as 

a relatively weak pathogen causing 10 

to 40% tree mortality. A more aggressive 

pathogen, causing nearly 100% mortality in affected elms, was 

discovered in the 1940’s in both North America and Europe 

(Brasier, 1990; 1991). This pathogen, O. novo-ulmi, was found to 

be two distinct subspecies, initially separated geographically 

(Brasier & Kirk, 2001). Ophiostoma novo-ulmi subsp. americana 

spread throughout North America in less than 40 years and 

was discovered in Europe in the 1960’s. Ophiostoma novo-ulmi 

subsp. novo-ulmi has seen similar spread in Eurasia (Brasier 

& Buck, 2001). The origin of Dutch elm disease is unknown. 

However, surveys in the Himalayas led to the discovery of a 

third species associated with Dutch elm disease: Ophiostoma 

himal-ulmi. This species, considered endemic to the Himalayas, 

is an aggressive pathogen of both North American and European 

elms (Brasier & Mehrotra, 1995). 

diseAse cycle 

The spread of Dutch elm disease is facilitated by elm bark 

beetles (Scolytus spp.), which preferentially breed in unhealthy 

trees. The Dutch elm disease fungus produces spores in the 

dead wood tissue surrounding the larval galleries. As these 

larvae develop into adults they become covered in spores, which 

can hitch a ride to other elm trees. As the adult beetles feed 

on the twigs and branches of healthy elm they create wounds 

and allow the spores to enter and spread through the vascular 

system of the tree (Pscheidt, 2011). Figure 285 shows the cycle 

of Dutch elm disease.

Elm.bark.beetles.transport. 

spores.to.new.hosts 44 . 


Fungal.spores.are.produced,.ready.. 

to.attach.to.emerging.adult.beetles 89 .. 

Figure 285: Cycle of Dutch elm disease. 

iMPActs 

.............The.fungus.infects.a.new.host.and. 

...............spreads.through.the.vascular.system 88 . 

..............Fungus.cuts.off.sap.transport 

............causing.wilting 70 . 

Elm is a popular ornamental tree commonly used to line 

urban roadsides. Dutch elm disease devastated the majority 

of these large picturesque elms (Fig. 286). Most trees die as a 

direct result of the disease or become weakened and succumb 

due to other environmental stresses. Elm mortality decreases 

forest biodiversity and wildlife habitat (NRCan, 2011). Economic 

impacts result from reductions in the quality and quantity of 

elm timber products as well as the negative effects associated 

with quarantine measures (CFIA, 2010). 

Figure 286: Comparing a road with healthy elm trees 41 (left) with one 

with elms affected by Dutch elm disease 90 (right). 

[ 237]

[ 238] 



The fungus that causes Dutch elm disease grows under the bark, 

along the vascular system of the tree, effectively blocking water 

transport from the roots to the leaves. The disease spreads via 

intermingling root systems or elm bark beetles, which can carry 

the spores. If the roots of neighbouring elm trees are close 

enough to each other, the fungus will spread from one tree to 

the other. Once in the roots the fungus can quickly spread as 

spores are carried through the vascular system (Haugen, 1998). 

There is one native and two exotic elm bark beetles that 

contribute to the long distance dispersal of Dutch elm disease 

(Fig. 287). See section 5.2.7 for detailed information on the 

biology and management of exotic elm bark beetles. These 

beetles complete part of their life cycle in the inner sapwood 

of elm trees. Beetles emerging from infected trees as adults 

carry fungal spores on their bodies to new healthy trees. They 

will often infest the upper branches and burrow into the wood 

to deposit their eggs and to feed. The spores can enter the tree 

through these wounds (Haugen, 1998). 

Figure 287: Banded elm bark beetle (Scolytus schevyrewi) 44 .




Prevention strategies for Dutch elm disease include destroying 

the breeding areas of insect vectors and eliminating other 

ways through which the disease can spread. Elm bark beetles 

breed in dead or dying elm material. It is best to burn or 

bury dead or infected branches. Elm firewood should not be 

left out in the open. If it cannot be burned or utilized right 

away it is best to completely encase the wood in plastic or 

tarps with the ends securely fastened or buried. Infected elm 

wood should be promptly disposed of (Davis & Meyers, 1997). 

Since Dutch elm disease can spread through root contact it is 

recommended to dig a trench around infected elm trees that 

are in close proximity to healthy elms. Ensure that the trench is 

deep enough to completely sever all connecting roots (Pscheidt, 

2011). 


Become familiar with the signs and symptoms of Dutch elm 

disease as well as those associated with elm bark beetles (See 

section 5.2.7). Look for signs of infected elm trees both in the 

woodlot and in the region. Contact a professional arborist to 

confirm the presence of Dutch elm disease and for help with 

management of infected trees (Davis & Meyers, 1997). 


Controlling Dutch elm disease involves removing any dead 

branches from apparently healthy trees or completely removing 

infected trees. Pruning dead branches from elm trees effectively 

removes any potential breeding grounds for elm bark beetles. 

This will greatly reduce the likelihood of infection. Pruning may 

also be employed to save infected elms if done early enough. 

Branches suspected of being infected should be debarked 

until no sapwood discolouration is observed. This is because 

discolouration is a result of infection and will aid in detecting 

how far the disease has spread. It is recommended to remove 

some of the healthy tissue below the infection. It is also very 

important to clean cutting tools between each use with rubbing 

alcohol to prevent spreading spores to other healthy trees or 

branches (Haugen, 1998). 

[ 239]

[ 240] 


It is good practice to remove elm trees that have been infected 

by Dutch elm disease. If sapwood discolouration has spread to 

the trunk this indicates that the tree can no longer be saved 

(Haugen, 1998). The presence of infected trees only contributes 

to the dispersal of Dutch elm disease. The resulting wood should 

be burned immediately. The stump should also be removed and 

burned. An alternative involves grinding the stump so that it is 

10cm below the soil line and then completely covering it with 

soil (Davis & Meyer, 1997). 

Fungicide injections have been used to control Dutch elm 

disease but they can be costly and problematic. These injections 

may wound the tree, causing discolouration and decay around 

the injection area (Shigo & Campana, 1977). In addition, Dutch 

elm disease seems to be developing a certain level of resistance 

to these fungicides, thereby decreasing their effectiveness. 

Fungicide applications are only feasible for use on high value 

ornamental trees. Application across an entire woodlot is 

considered impractical. Management activities that promote 

diverse and healthy woodlots are the best means of control for 

Dutch elm disease (Hubbes, 1999).


5.2.7 

elm Bark Beetles 

(Scolytus multistriatus & S. schevyrewi) 


lesser european elm bark beetle, smaller european elm bark beetle, 

Banded elm bark beetle 


Figure 288: Elm bark beetles 44 . 

241 

[ ]

[ 242] 


orGAnisMs 


There are two exotic elm bark beetles in Ontario: the banded 

elm bark beetle (Scolytus schevyrewi) and the European elm bark 

beetle (S. multistriatus) (Fig. 289). The banded elm bark beetle is 

brown with a dark band across its wing covers. It is generally 3 

to 4mm long. The European elm bark beetle lacks the band and 

does not exceed 3mm in length. Only the European elm bark 

beetle has a spine on the lower abdomen (Lee et al. 2006). 

Figure 289: Comparing the banded elm bark beetle 44 (left) and the 

European elm bark beetle 31 (right). 

host trees 

The European and the banded 

elm bark beetle are both 

host specific, feeding and 

breeding on elm trees (Ulmus 

sp.). Three native species of 

elm occur in Ontario: white 

elm (U. americana), slippery 

elm (U. rubra), and rock 

elm (U. thomasii). See figure 

280 (section on Dutch elm 

disease) for a comparison of 

leaf morphologies and native 

range maps for these species. 

Elm is a popular ornamental 

tree planted along roadsides 

and parks (Fig. 290). Many 

are exotic. Exotic elms found 

in Ontario include wych elm 

(U. glabra), English elm (U. 

procera) and Siberian elm (U. 

pumila); all are susceptible to 

European or banded elm bark 

beetles (Sargent et al. 2008; 

Farrar, 1995). 

Figure 290: Elm is a popular ornamental tree 41 .

5.2.7 ELM BARK BEETLES (Scolytus multistriatus & S. schevyrewi) 


exit & entrAnce holes 

Female European and banded elm bark beetles create an 

entrance hole and lay their eggs in the bark of elm trees. When 

the eggs hatch, the larvae develop inside the tree and emerge 

as adults through 2mm circular exit holes in the trunk or 

branches (Sargent et al. 2008) (Fig. 291). 

Figure 291: Exit holes created by the European elm bark beetle 92 (left) and the banded elm bark beetle 44 (right). 

lArvAl GAlleries 

Female European and banded elm bark beetles create and lay 

their eggs in a vertical tunnel under the bark. Larvae make 

tunnels in an outward fashion, creating a fan-like appearance 

on both sides of the egg gallery (Fig. 292). Developing and 

feeding larvae create sawdust that can be seen in the larval 

tunnels and exit holes (Lee et al. 2006) (Fig. 293). 

Figure 292: Larval galleries created by the European elm bark beetle 57 . 

[ 243]

[ 244] 


Figure 293: Sawdust created by banded elm bark beetle larvae 44 . 


syMPtoMs 

native elm Bark Beetle (Hylurogoplinus rufipes) 

The native elm bark beetle can be distinguished from exotic 

elm bark beetles by its metallic gold colour (Fig. 294). Adult 

females create egg galleries by tunneling horizontally across 

the wood grain, whereas exotic elm bark beetles create vertical 

tunnels (Fig. 295). Native elm bark beetles leave behind 

V-shaped markings beside their exit holes, a characteristic not 

seen for exotic elm bark beetles (CFS, 2011). 

Figure 294: Comparing native 92 (left) and European elm bark beetles 92 (right).

Figure 295: Comparing larval galleries of the native elm bark beetle 64 (left) with those of the European elm 

bark beetle 41 (right). 


European Elm Bark Beetle (S. multistriatus) 





........Family. Curculionidea 

..........Genus. Scolytus 


Species Scolytus multistriatus 

BioloGy 


Banded Elm Bark Beetle (S. schevrewi) 





........Family. Curculionidea 

..........Genus Scolytus 

Species Scolytus schevrewi 

As its common name indicates, the European elm bark beetle is 

native to Europe. First found in Massachusetts in 1909 (Lee et 

al. 2009), it has since spread throughout Canada (NRCan, 2011). 

The banded elm bark beetle is native to Asia and is a relatively 

recent introduction to North America. It was first observed 

in 2003 in both Colorado and Utah but evidence from past 

collections indicates that it has been in North America since 

[ 245]

[ 246] 


the 1990’s (Negrón et al. 2005). In Canada, it was first detected 

in Alberta in 2006 with subsequent introductions in Manitoba, 

Ontario, Saskatchewan (Langor et al. 2009), and, most recently, 

British Columbia (Humble et al. 2010). 

liFe cycle 

Elm bark beetles overwinter as adults under the bark of elm 

trees. They emerge in the spring and begin feeding on the inner 

tissues of small twigs and branches. It is at this stage that the 

elm bark beetle can transmit Dutch elm disease to new hosts 

(Ohmart, 1989). After feeding, the adult beetles seek out weak or 

dying elm trees to breed, which can become infected with Dutch 

elm disease (Rudinsky, 1962). Fallen elm logs with intact bark 

also attract breeding adults. Females create tunnel-like galleries 

in which they lay their eggs. Eggs hatch in the spring and the 

larvae feed and pupate for approximately 2 months before 

emerging as adults (Ascerno & Wawrzynski, 1994) (Fig. 296). 

Adult.feeding.on.a.healthy.. 

elm 44 

Adults.emerge 44 

Figure 296: Life-cycle of elm bark beetles. 

iMPActs 

.............Breeding.on.dead.or.stressed 

...............elm 44 

..............Larval.development.in.dead 

............or.stressed.elm 44 . 

Elm bark beetles alone appear to have minimal ecological impact. 

Although the larval tunnels under the bark may affect lumber 

value, considering that elm bark beetles usually breed in dead 

or dying elm trees, the risk to lumber value is low. Their greatest 

impact lies in their role as vectors of Dutch elm disease, a deadly 

condition that affects the tree’s vascular system (Seybold et al. 

2008). See section 5.2.6 for information on Dutch elm disease.



Elm bark beetles were most likely introduced in North America 

via wood packaging materials. Any materials with intact bark 

are capable of carrying elm bark beetles. Dispersal is likely 

facilitated by the transportation of nursery stock and elm wood 

products such as firewood. Adult beetles are not strong flyers. 

Instead, wind currents may be more important factors in their 

natural dispersal (Sargent et al. 2008). 



Adult elm bark beetles are attracted to wounded elm trees. 

Destroy all dead or dying elm trees and elm wood by chipping, 

burning or burying. This will decrease the number of 

potential breeding sites available for elm bark beetles. Do not 

use or transport elm firewood. Elm bark beetles are capable 

of completing their life cycle on elm logs provided there is a 

small patch of bark left intact (Sargent et al. 2008). If pruning 

activities are scheduled it is best to wait until the late fall or 

winter when adult beetles are less active (Byers et al. 1980). The 

fungal agent of Dutch elm disease has been shown to release 

chemical signals that attract elm bark 

beetles (McLeod et al. 2005). Look for 

signs and symptoms of this disease 

and consider eliminating infected trees. 

Refer to section 5.2.6 for management 

strategies for Dutch elm disease. 

eArly detection 

techniques 

Regularly look for signs and symptoms of 

elm bark beetle establishment, including 

yellowing or wilting foliage, circular exit 

holes in the bark, and declining elm tree 

health. When signs or symptoms of the 

beetle are found peeling the bark away 

to determine whether the characteristic 

larval galleries are present can aid in 

identification (Sargent et al. 2008) (Fig. 

297). 

Figure 297: Larval galleries of 

European elm bark beetle 94. 

[ 247]

[ 248] 



Dead elm trees, logs and branches should be destroyed through 

chipping, burning or burying. This will decrease the number 

of available breeding sites and prevent populations of elm bark 

beetles from increasing (Sargent et al. 2008). Keep elm trees 

healthy and properly pruned. However, only prune during the 

late fall and winter when adult beetles are less active (Byers et 

al. 1980). 

Woodlot management activities should be geared towards 

increasing tree health because elm bark beetles are attracted 

to stressed trees. This can be done by increasing tree species 

diversity, thinning to reduce competition and removing 

damaged or diseased trees (Seybold et al. 2008). Elm bark 

beetles will also invade freshly cut logs, recently fallen trees, 

and those affected by fire. Thus, it is important to monitor 

elm trees in the woodlot and eliminate any potential breeding 

grounds (Rudinsky, 1962). 

Since elm bark beetles spend the majority of their life cycle 

within the tree, insecticides are generally applied to the outer 

bark only as a preventative measure. This is because these 

chemicals are only effective if they come into direct contact 

with the adult beetles. Consequently, insecticide use may be 

impractical (Seybold et al. 2008).


5.2.8 

Butternut canker 

(Ophiognomonia clavigignenti-juglandacearum) 


Butternut disease 


Figure 298: Butternut Canker 41 . 

249 

[ ]

[ 250] 


cAusAl orGAnisM 


Butternut (Juglans cinerea) is native to Ontario and the 

primary host of the butternut canker fungus (Ophiognomonia 

clavigignenti-juglandacearum). Previously classified as 

Sirococcus, this fungus has recently been reclassified to the 

genus Ophiognomonia (Broders & Boland, 2011). Species in the 

Ophiognomonia genus are characterized by their elongated 

ascospores that infect a variety of plant families, including 

Juglanaceae to which butternut belongs (Walker et al. 2012). 

Ophiognomonia claviginenti-juglandacearum reproduces 

asexually. Fungal spores enter butternut trees through wounds 

or cracks in the bark. The fungus forms projecting structures 

under the bark called hyphal pegs (Fig. 299). These hyphal pegs 

cause the bark to crack and split open. Spores are released into 

the environment through these openings (Nair et al. 1979). 

Figure 299: Hyphal pegs of Ophiognomonia clavigignentijuglandacearum 

95 . 

host trees 

Butternut (Juglans cinerea) is the main host of O. clavigignentijuglandacearum 

(Fig. 300). A native to North America, this 

tree species generally occurs in isolated patches in Ontario, 

Quebec and New Brunswick (NRCan, 2011). Butternut has been 

federally and provincially listed as an endangered species due 

to the devastating impacts of O. clavigignenti-juglandacearum 

(Environment Canada, 2011; OMNR, 2012b). Other members 

of the walnut family have shown some susceptibility to the 

disease but, unlike butternut, do not appear to be killed by it 

(Ostry et al. 1996).

5.2.8 BUTTERNUT CANKER (Ophiognomonia clavigignenti-juglandacearum) 

cAnkers 

Figure 300: Butternut leaves 5 (left) and fruit 5 (right). 


The main symptom of O. clavigignenti-juglandacearum infection 

is the formation of stem and branch cankers. These cankers 

appear as large black spots on the branches and trunks of 

butternut trees (Fig. 301). Cankers are black and often release 

a black sappy substance in the spring. In the summer they dry 

out, having a sooty black appearance with shades of white along 

the outer edges (Davis & Meyer, 1997). 

Figure 301: Butternut cankers in the spring 41 (left) and summer 41 (right). 

[ 251]

[ 252] 


sAPwood discolourAtion 

Removing the bark in areas where a suspected canker is forming 

will reveal a black, darkened area (Fig. 302). As cankers form and 

the infection spreads, the sapwood underneath the bark decays 

and turns black. The fungus will continue to spread and, as the 

disease intensifies, large areas of dead and blackened cambium 

tissue appear (Ostry et al. 1994). 

MortAlity 

As infection progresses and decay continues within the 

cambium layer, branches become girdled and die. During 

heavy rainfall, spores are carried downward and establish new 

infections in multiple areas of the main trunk. The numerous 

cankers formed on the main trunk eventually kill the tree 

(Fig. 303). Epicormic branching is common on stressed trees. 

However, these branches quickly become infected and die 

(Davis & Meyer, 1997). 

Figure 302: Sapwood 

discolouration from dead 

cambium tissue 96 . 

Figure 303: Butternut killed 

as a result of O. clavigignentijuglandacearum 

infection 69 . 


syMPtoMs 

Butternut trees form similar cankers and experience crown 

dieback as a result of agents other than O. clavigignentijuglandacearum. 

A similar fungus, Melanconis oblongum, 

which can co-occur with O. clavigignenti-juglandacearum


infects dead butternut tissue. However, it is not known to cause 

butternut mortality (Ostry et al. 1994). Bacterial blight caused 

by Xanthomonas arboricola pv. juglandis infection also causes 

cankers in the form of black lesions but branch and crown 

dieback does not generally occur as a result. Bunch disease, 

caused by unknown bacteria, does not produce cankers but 

has been shown to cause increased lateral growth that may 

be mistaken for the epicormic shoots produced in response 

to butternut canker (Pijut, 2006). Cankers can also form from 

physical injury or frost damage (Harrison & Hurley, 2001). It 

is recommended to consult a professional to help identify the 

cause of butternut decline (Fig. 304). 


Figure 304: Butternut affected by butternut canker 97 . 

BioloGy 



The origin of O. clavigignentijuglandacearum 

is unknown. The 


fungus was first observed in North 


America in 1967 but it was not until 

1979 that it was identified as the 

......Order. Diaporthales 

causative agent of butternut canker. 

........Family. Gnomoniaceae 

Today it is found in the eastern United 

States and southeastern Canada 

..........Genus. Ophiognomonia 

where it has spread throughout 

the entire range of its host species 

Species Ophiognomonia clavigignentijuglandacearum 

(Schlarbaum et al. 1997; Harrison 

& Hurley, 2001) (Fig. 305). It was 

confirmed in Ontario in 1991 from 

a sample collected in Ipperwash Provincial Park, south of Lake 

Huron (Davis et al. 1992). 

[ 253]

[ 254] 


Figure 305: Native range of butternut in Ontario 64 . 

diseAse cycle 

Spores of O. clavigignenti-juglandacearum infect butternut 

trees through buds, leaf scars and other wounds found in the 

bark. The fungal infection causes cankers under which hyphal 

pegs exert pressure on the inner bark, causing it to split. Fungal 

spores can then escape through these cracks and be carried 

by water, wind or other organisms such as insects, birds and 

rodents, to new butternut hosts (Nair et al. 1979). Heavy rainstorms 

can accelerate the infection in individual butternut trees 

because the water runs down the trunk collecting spores and 

transporting them downward. As such, multiple cankers can 

form on the same tree (Tisserat & Kuntz, 1983). The fungus has 

been reported to survive for up to 2 years on dead trees (Ostry, 

1996). Figure 306 shows the cycle of butternut canker. 

Hyphal.pegs.rupture.the.bark. 

and.spores.are.released. 95 

Establishes.itself.under.the.. 

bark.and.forms.fruiting.bodies 41 

.............Spores.are.transported.by.wind, 

..............rain.or.biota.to.new.host.trees 52 

..............Spores.infect.through.buds,.leaf 

...........scars.and.wounds.in.the.bark 5 . 

Figure 306: Cycle of butternut canker.


iMPActs 

The introduction of butternut canker has had devastating 

impacts on butternut populations and forest biodiversity. The 

fungal infection can kill trees of all ages and sizes (Innes et 

al. 2001). The majority of infected trees die rapidly after initial 

infection. Both the above and below-ground portions of the 

tree die as a result of the disease (Environment Canada, 2010). 

Butternut produces a seed crop that is a highly valued food 

source for wildlife such as small mammals and birds (Rink, 

1990). Humans can also benefit from the commercial value of 

the fruits. In addition, the lumber is considered to be of high 

value when unaffected by the disease for its uses in woodworking 

(Ostry & Pijut, 2000). 


The butternut canker fungus can spread within the host tree. 

Externally, as water runs down from the top branches it collects 

spores from existing cankers and transports them to lower 

wounds (Tisserat & Kuntz, 1983). Spores have also been shown 

to travel up to 45m away from their original host via wind 

currents. Insects such as beetles can carry the spores on their 

bodies if they come into physical contact with cankers. It has also 

been speculated that birds play a role in the spread of butternut 

canker (Halik & Bergdahl, 2002). Humans also contribute to the 

spread of butternut canker through the movement of firewood, 

seeds and nursery stock (Ostry & Woeste, 2004). 



There is a lack of known preventative measures against butternut 

canker other than practicing appropriate woodlot management 

practices and keeping the forest healthy. Butternut canker 

can easily infect butternut trees through wounds in the bark. 

Minimizing physical damage during management activities 

is recommended. Healthy trees are generally less susceptible 

to insect attack and disease than stressed or weakened trees. 

Keeping a diverse and healthy woodlot could help to prevent 

the introduction of pathogens such as the causing agent of 

butternut canker (Ostry et al, 1996). 

[ 255]

[ 256] 



Monitor butternut trees in the woodlot and look for signs 

and symptoms of the disease. It is prohibited to cut down a 

naturally occurring butternut tree as the species is protected 

under Ontario’s Endangered Species Act. Woodlot owners 

considering removal of butternut should contact the OMNR, 

even if the tree(s) is/are infected (OMNR, 2011). 


As a woodlot owner, the best management strategy to prevent 

butternut canker is to promote a healthy woodlot. When 

performing woodlot management or timber harvest activities 

it is best to remove groups of trees that may be shading out 

butternut. Butternut is a shade intolerant species and requires 

canopy gaps to promote regeneration and healthy growth. 

Managing both healthy and infected butternut trees is an 

important goal in the conservation of this species. Infected 

butternut trees may develop a certain level of resistance, which 

may be beneficial in future breeding programs (DesRochers, 

2009). Butternut trees showing little to no signs of infection 

may possess a certain level of natural resistance to the disease. 

Woodlot owners are encouraged to report these trees to the 

Forest Gene Conservation Association. This organization has 

been recording the locations of potentially resistant butternut 

trees for possible use in breeding programs. These programs 

consist of crossing resistant individuals with the objective of 

creating a disease-resistant population (FGCA, n.d.). 

Federal law currently protects butternut. Given its status, even 

heavily infected trees should be treated, not cut down. It is an 

offense to possess or sell naturally occurring butternut wood 

under the Endangered Species Act, 2007. In fact, a woodlot owner 

can only remove a butternut tree if considered non-retainable. 

That is, if infection is so severe that it is no longer possible 

to save the tree. An expert accredited as a Butternut Health 

Assessor must identify the tree as non-retainable. Woodlot 

owners can find a Butternut Health Assessor by contacting the 

OMNR (OMNR, 2011).


5.2.9 

dogwood Anthracnose 

(Discula destructiva) 


dogwood disease 



Dogwood anthracnose (Discula destructiva) is an airborne 

fungal disease of flowering dogwood (Cornus florida) and 

Pacific dogwood (C. nuttallii) (Hibben, 1990). The origin of D. 

destructiva is unknown (Zhang & Blackwell, 2001). Black or 

brown spots bordered in purple on the leaves are indicative of 

dogwood anthracnose (Fig. 307). These spots eventually turn 

into holes. These symptoms first appear in the lower crown and 

progress upwards. Infection on the leaves can spread through 

the petioles into the twigs. Cankers begin to appear and can 

eventually girdle and kill the tree (Sinclair & Lyon, 2005). 

Figure 307: Leaf lesions 83 (left) and cankers 83 (right) caused by dogwood 

anthracnose. 

[ 257]

[ 258] 


iMPActs 

An abundance of white showy flowers and 

bright red berries contribute to flowering 

dogwood’s popularity as an ornamental 

tree (Fig. 308). Its fruit and seeds have a 

high nutritional content that makes them an 

invaluable food source for wildlife (Rossell 

et al. 2001). 

Dogwood anthracnose has had devastating 

effects on flowering dogwood in Ontario with 

an estimated decline of 7-8% per year. This 

decline has led to the provincial and federal 

governments listing flowering dogwood 

as an endangered species (Bickerton & 

Thompson-Black, 2010). 

control 

Flowering dogwood is an endangered species protected by 

the Endangered Species Act, 2007. In Canada, the native range 

of flowering dogwood is the Carolinian forest of southern 

Ontario (Bickerton & Thompson-Black, 2010)(Fig. 309). A healthy 

flowering dogwood is less susceptible to infection than a 

weakened or stressed tree. As such, dead or infected branches 

should be pruned, removed and destroyed. Clean cutting tools 

with rubbing alcohol between each use to prevent spreading the 

disease (Pecknold et al. 2001). Report any potentially resistant 

trees to a species-at-risk biologist at the OMNR. Potentially 

resistant trees may be useful for research programs investigating 

ways of saving flowering dogwood populations and controlling 

dogwood anthracnose (Bickerton & Thompson-Black, 2010). 

Figure 309: Native range of flowering dogwood (Cornus florida) 64 . 

Figure 308: Showy flowers 15 (top) 

and bright red berries 66 (bottom) of 

flowering dogwood.



Thousand cankers disease is caused by a fungal pathogen 

(Geosmithia morbida) and dispersed by the native walnut twig 

beetle (Pityophthorus juglandis) (Fig. 310). The disease mainly 

affects black walnut (Juglans nigra). Yellowing of the leaves is 

the first sign of infection followed by dying branches. Cankers 

form around entry wounds created by the beetle (Fig. 311). Trees 

usually die within three years as a result of the disruption of 

nutrient transport (Leslie et al. 2010). The disease received its 

name for the numerous cankers that form on the trunk and 

branches of infected trees (Cranshaw & Tisserat, 2008). 

Figure 310: Walnut twig beetle 

(Pityophthorus juglandis) 99. 

5.2.10 

thousand cankers disease 

(Geosmithia morbida) 


tcd 


Figure 311: Cankers caused by thousand cankers disease on a walnut 

sapling 100 (left) and in the sapwood of a mature walnut tree 101 (right). 

[ 259]

[ 260] 


iMPActs 

Thousand cankers disease can wipe out entire native populations 

of black walnut (Fig. 312). Until recently, the disease was confined 

to western North America where it was causing widespread 

mortality of ornamental black walnut. It has recently been 

detected in Tennessee (2010), Virginia (2011) and Pennsylvania 

(2011), which is a major problem as this comprises the native 

range of black walnut (Grant et al. 2011). Black walnut is an 

economically important species that is harvested for both its 

lumber and fruit (MacDaniels & Lieberman, 1979). They produce 

a nut crop rich in protein, which is an important energy source 

for wildlife (Smith & Follmer, 1972). 

Figure 312: Native range of black walnut (Juglans nigra) 64 . 

control 

Thousand cankers disease is not present in Ontario. However, it 

is important to be prepared in the event that the disease complex 

finds its way into Canada. Monitor the health of black walnut trees 

on the property and look for signs and symptoms of disease. If 

you suspect that you have a black walnut tree 

infected with thousand cankers disease contact 

the CFIA and/or the OMNR. In areas infested 

by thousand cankers disease, infested walnut 

trees should be removed to eliminate sources 

of infection (Fig. 313). Walnut wood should be 

heat-treated and the bark should be removed. 

Walnut twig beetles complete their life cycle 

under the bark. Therefore, logs without bark will 

not support these insect vectors. Contaminated 

wood should never be moved outside of infested 

areas (Cranshaw & Tisserat, 2008). 

Figure 313: Controlling thousand 

cankers disease 100 .


5.2.11 

Pear thrips 

(Taeniothrips inconsequens) 


Fruit tree thrips 



Pear thrips (Taeniothrips inconsequens) are small, black insects 

that are approximately 1.5mm in length (Fig. 314). They spend 

part of their life cycle in the soil and part on hardwood trees. 

In the spring, adults emerge from the soil and enter tree buds. 

They feed on immature leaves and lay their eggs within leaf 

tissues. Larvae are clear and very hard to see. They eventually 

drop to the soil where they pupate and emerge as adults the 

following year (Hoover, 2002). 

Figure 314: Pear thrips in the adult 40 (left) and larva 40 (right) life stages. 

[ 261]

[ 262] 


iMPActs 

Pear thrips were introduced to North America from Eurasia in 

the early 1900’s (Palm & Gardescu, 2008). Sugar maple seems to 

be the preferred host for this exotic insect, although damage 

has been observed on a variety of other hardwoods such as 

beech, ash, serviceberry and cherry as well as on a variety of 

herbaceous plants commonly found in forest understories (Palm 

& Gardescu, 2008). The feeding adults cause leaves to crinkle and 

wilt (Fig. 315). Crown defoliation and yellowing is often a sign 

of a high density of pear 

thrips. Seed production 

and sap yields may also 

be adversely affected, 

which in turn could 

have negative effects 

on regeneration and 

future forest diversity. 

Hardwoods affected by 

pear thrips defoliation 

may be more susceptible 

to other insect pests 

and pathogens (Hoover, 

2002). Gardescu (2003) 

found that pear thrips 

were the primary cause 

of hardwood seedling 

mortality in a sugar 

maple forest in New York. 

Figure 315: Damage to maple leaves caused by pear 

thrips 102 . 

control 

Pear thrips can be very difficult to control when populations 

reach high densities. Insecticides are not very effective because 

the adults spend most of the time within the leaf buds and 

tissues. Furthermore, insecticides should not be applied to 

sugar maples in woodlots utilized for maple syrup production. 

In years of severe defoliation, maple syrup producers may have 

to avoid tapping infected trees to counteract the effect of the 

insects and prevent maple mortality (Hoover, 2002). 

Several native insects and fungal pathogens affect pear thrips 

in North America but more research is needed to determine 

whether they can be augmented to provide any level of 

biological control. However, heavy infestations seldom last for 

more than one year (Palm & Gardescu, 2008). More research on 

the potential long-term consequences of this invasive species 

is necessary.

6.0 

APPendices 

Appendix 1: Invasive plants considered a high priority for management based on their associated risks. 

Species Risk.Category Risk References 

Norway.maple . 


Tree-of-heaven. 

(Ailanthus 

altissima) 

Garlic.mustard. 


Economic 

Environmental 

Cause.reduced.hardwood.growth Galbraith-Kent.&.Handel,.2012 

Suppress.hardwood.regeneration 

Cause.the.loss.of.biodiversity 

Fang.&.Wang,.2011 

Galbraith-Kent.&.Handel,.2008 

Martin,.1999 

Reinhart.et.al..2005;.2006 

Wycoff.&.Webb,.1996 

Bertin.et.al..2005 

.Fang,.2005 

.Reinhart.et.al..2005 

.Wyckoff.&.Webb,.1996 

Affect.ecosystem.function Gómez-Aparicio.&.Canham,.2008a 

Economic Suppress.hardwood.regeneration 

Environmental 

Cause.the.loss.of.biodiversity 

Heisey,.1990 

Lawrence.et.al..1991 

Mergen,.1959 

Knapp.&.Canham,.2000 

Small.et.al..2010 

Affect.ecosystem.function Gómez-Aparicio.&.Canham,.2008b 

Social Impacts.on.human.health 

Economic 

Environmental 

Ballero.et.al..2003 

Derrick.&.Darley,.1994 

Cause.reduced.hardwood.growth Stinson.et.al..2006 


Meekins.&.McCarthy,.1999 

Stinson.et.al..2006 

Cause.the.loss.of.biodiversity Stinson.et.al..2007 

Affect.ecosystem.function 

Harm.species-at-risk 

Burke,.2008 

Cantor.et.al..2011 

Koch.et.al..2011 

Rodgers.et.al..2008 

Renwick,.2002 

Porter,.1994 

6.0 

[ 263]

[ 264] 



Barberry.. 

(Berberis spp.) 

English.Ivy. 


Himalayan. 

balsam. 

(Impatiens 

glandulifera) 

Japanese. 

knotweed 

.(Fallopia 

japonica) 

Common. 

buckthorn. 

(Rhamnus 

cathartica) 

Periwinkle.. 

(Vinca minor) 

Dogstrangling.vine. 

(Vincetoxicum 

spp.) 

Economic Cause.reduced.hardwood.growth Silander.Jr..&.Klepeis,.1999 

Environmental 

Cause.the.loss.of.biodiversity Kourtev.et.al..1998 

Affect.ecosystem.function 

Social Impacts.on.human.health 

Economic 

Cause.hardwood.mortality 


Environmental Cause.the.loss.of.biodiversity 

Social 

Appendix 1: cont’d. 

Impacts.on.human.health 

Affect.recreational.enjoyment 

Ehrenfeld.et.al..2001 

Kourtev.et.al..1998;.2002a;.2002b;.2003 

Elias.et.al..2006 

Williams.et.al..2009 

Schnitzer.&.Bongers,.2002 

Swearingen.&.Diedrich,.2006 

Thomas,.1980 

Dlugosch,.2005 

Schnitzer.&.Bongers,.2002 

Dlugosch,.2005 

Harmer.et.al..2001 

Thomas,.1980 

Swearingen.et.al..2010 

Economic Suppress.hardwood.regeneration Maule.et.al..2000 

Pyšek.&.Prach,.1995 





Environmental Cause.loss.of.biodiversity 

Environmental Affect.ecosystem.function 

Andrews.et.al..2009 

Clements.et.al..2008 

Perrins.et.al..1993 

Beerling.et.al..1994 

Weber,.2003 

Aguilera.et.al..2010 

Beerling.et.al..1994 

Maerz.et.al..2005 

Delanoy.&.Archibold,.2007 

Mascaro.&.Schnitzer,.2007 

Klionsky.et.al..2010 

Knight.et.al..2007 

Heneghan.et.al..2004;.2006;.2007 

Knight.et.al..2007 

Economic Suppress.hardwood.regeneration Darcy.&.Burkart,.2002 

Environmental Cause.loss.of.biodiversity Drake.et.al..2003 

Environmental Affect.ecosystem.function Bultman.&.DeWitt,.2008 

Economic Suppress.hardwood.regeneration DiTommaso.et.al..2005 

Environmental Cause.loss.of.biodiversity 

Cappuccino,.2004 

Kricsfalusy.&.Miller,.2010 

Environmental Affect.ecosystem.function Smith.et.al..2008 

Environmental Harm.a.species-at-risk 

DiTommaso.&.Losey,.2003 

Ladd.&.Cappuccino,.2005

Appendix 2: Invasive insects and pathogens considered a high priority for management based on their 

associated risks. 


Emerald.ash.borer. 


Asian.long-horned. 

beetle.(Anoplophora 

glabripennis) 

Chestnut.Blight. 

(Cryphonectria 

parasitica) 

Gypsy.moth. 


APPENDICES 

Economic Cause.hardwood.mortality 

Environmental. Cause.loss.of.biodiversity 

Cappaert.et.al..2005 

MacFarlane.&.Meyer,.2005 

Poland.&.McCullough,.2006 

Raupp.et.al..2006 

Gandhi.&.Herms,.2010 

Kimoto.&.Duthie-Holt,.2006 

Poland.&.McCullough,.2006;. 

Social Interfere.with.a.traditional.lifestyle Herms.et.al..2004 

Social Reduce.aesthetic.values.of.the.forest 

Cartwell,.2007 

Economic Cause.reduced.hardwood.growth Dodds.&.Orwig,.2011 

Economic. Cause.hardwood.mortality 


Hu.et.al..2009 

Raupp.et.al..2006 

Environmental. Cause.loss.of.biodiversity Kimoto.&.Duthie-Holt,.2006 

Social Interfere.with.a.traditional.lifestyle Cartwell,.2007 

Social. Reduce.aesthetic.values.of.the.forest 

Economic Cause.reduced.hardwood.growth 



Nowak.et.al..2001 

McEwan.et.al..2006 

Paillet,.2002 

Tindall.et.al..2004 

Griffin,.2000 

Sinclair.&.Lyon,.2005 

Environmental. Cause.loss.of.biodiversity Sinclair.&.Lyon,.2005 

Environmental. Harm.a.species-at-risk 

OMNR,.2012b 

.Sinclair.&.Lyon,.2005 

Social Interfere.with.a.traditional.lifestyle Youngs,.2000 

Social Reduce.aesthetic.values.of.the.forest Anagnostakis,.1982;.1987 

Economic. Cause.reduced.hardwood.growth 



Social. Affect.recreational.enjoyment 

Baker,.1941 

Tobin.&.Liebold,.2011 

Baker,.1941 

Davidson.et.al..1999 

Fajvan.&.Wood,.1996 

Tobin.&.Liebold,.2011 

Hollenhorst.et.al..1991 

Régnière.et.al..2009 

Leuschner.et.al..1996 

Régnière.et.al..2009 

Social. Impacts.on..human.health McManus.et.al..1989 

6.0 

[ 265]

[ 266] 



Beech.bark.disease. 

(Neonectria faginata) 

Dutch.elm.disease. 


Elm.bark.beetles. 

(Scolytus spp.) 

Butternut.canker. 

(Ophiognomonia 

clavigignentijuglandacearum) 

Appendix 2: Cont’d. 



Houston,.1994 

McCullough.et.al..2005 

Shigo,.1972 

Hane,.2003 

Nyland.et.al..2006 

Environmental. Cause.loss.of.biodiversity McCullough.et.al..2005 

Environmental. Affect.ecosystem.function 

McCullough.et.al..2005 

Storer.et.al..2005 

Social Reduce.aesthetic.values.of.the.forest Sinclair.&.Lyon,.2005 

Social. Affect.recreational.enjoyment Heyd,.2005 

Economic. Cause.hardwood.mortality Gibes,.1978 

Environmental. Cause.loss.of.biodiversity NRCan,.2011 



Davis.&.Meyer,.1997 

Sinclair.&.Lyon,.2005 

Negrón.et.al..2005 

Seybold.et.al..2008 

Environmental. Cause.loss.of.biodiversity NRCan,.2011 

Economic Cause.hardwood.mortality Davis.&.Meyer,.1997 

Environmental. Cause.loss.of.biodiversity 


Schlarbaum.et.al..1997 

Environmental. Harm.a.species-at-risk OMNR,.2012b 



Ostry.et.al..1994

7.0 

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Anulewicz,.A..C.,.McCullough,.D..G.,.&.Cappaert,.D..L..(2007).. 

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2.. James.Smedley,.James.Smedley.Outdoors,.www. 

jamessmedleyoutdoors.com 

3.. Jeff.&.JoAnn.St..Pierre,.North.Country.Photography,. 

www.jeffnorthcountry.com 

4.. Jan.Samanek,.State.Phytosanitary.Administration,. 

Bugwood.org 

5.. Paul.Wray,.Iowa.State.University,.Bugwood.org 

6.. Leslie.J..Mehrhoff,.University.of.Connecticut,.Bugwood. 

org 

7.. Bill.Cook,.Michigan.State.University,.Bugwood.org 

8.. Keith.Kanoti,.Maine.Forest.Service,.Bugwood.org 

9.. John.Ruter,.University.of.Georgia,.Bugwood.org 

10.. The.Dow.Gardens.Archive,.Dow.Gardens,.Bugwood.org 

11.. Chuck.Bargeron,.University.of.Georgia,.Bugwood.org 

12.. James.H..Miller,.USDA.Forest.Service,.Bugwood.org 

13.. Jody.Shimp,.Illinois.Department.of.Natural.Resources,. 

Bugwood.org 

14.. Walter.Muma,.Ontario.Trees.and.Shrubs,.http:// 

ontariotrees.com 

15.. Chris.Evans,.Illinois.Wildlife.Action.Plan,.Bugwood.org 

16.. Karan.A..Rawlins,.University.of.Georgia,.Bugwood.org 

17.. Dave.Powell,.USDA.Forest.Service,.Bugwood.org 

18.. Gyorgy.Csoka,.Hungary.Forest.Research.Institute,. 

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20.. Walter.Muma,.Ontario.Wildflowers,.http:// 

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8.0 

PhotoGrAPhy credits 

21.. Allen.Bridgman,.South.Carolina.Department.of.Natural. 

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22.. Steve.Manning,.Invasive.Plant.Control,.Bugwood.org 

23.. Mary.Burrows,.Montana.State.University,.Bugwood.org 

24.. Donna.R..Ellis,.University.of.Connecticut,.Bugwood.org 

25.. Forest.&.Kim.Starr,.Starr.Environmental,.Bugwood.org 

26.. Jil.Swearingen,.USDI.National.Park.Service,.Bugwood.org 

27.. Charles.T..Bryson,.USDA.Agricultural.Research.Service,. 

Bugwood.org 

28.. Deborah.L..Miller,.USDA.Forest.Service,.Bugwood.org 

29.. Thaddeus.Lewandowski,.Algoma.University.Alumni,. 

Sault.Ste..Marie 

30.. Louisiana.State.University.AgCenter.Archive,.Louisiana. 

State.University.AgCenter,.Bugwood.org 

31.. Thérèse.Arcand,.Natural.Resources.Canada,.Canadian. 

Forest.Service,.Laurentian.Forestry.Centre. 

32.. Robert.H..Mohlenbrock.@.USDA-NRCS.PLANTS/USDA. 

NRCS..1995..Northeast.wetland.flora:.Field.office.guide. 

to.plant.species..Northeast.National.Technical.Center,. 

Chester. 

33.. Bruce.Moltzan,.USDA.Forest.Service,.Bugwood.org 

34.. Vladimir.Petko,.V.N..Sukachev.Institute.of.Forest.SB.RAS,. 

Bugwood.org 

35.. Nancy.Loewenstein,.Auburn.University,.Bugwood.org 

36.. James.H..Miller.&.Ted.Bodner,.Southern.Weed.Science. 

Society,.Bugwood.org 

37.. Peggy.Greb,.USDA.Agricultural.Research.Service,. 

Bugwood.org 

38.. Erich.G..Valley,.USDA.Forest.Service.–.SRS-4552,. 

Bugwood.org 

39.. David.Cappaert,.Michigan.State.University,.Bugwood.org 

40.. Pennsylvania.Department.of.Conservation.and.Natural. 

Resources.–.Forestry.Archive,.Bugwood.org 

41.. Joseph.O’Brien,.USDA.Forest.Service,.Bugwood.org 

42.. Jared.Spokowsky,.New.York.State.Department.of. 

Agriculture.and.Markets,.Bugwood.org 

43.. Daniel.Herms,.The.Ohio.State.University,.Bugwood.org 

44.. Whitney.Cranshaw,.Colorado.State.University,.Bugwood. 

org 

45.. William.Jacobi,.Colorado.State.University,.Bugwood.org 

46.. Howard.Ensign.Evans,.Colorado.State.University,. 

Bugwood.org 

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47.. James.A..Copony,.Virginia.Department.of.Forestry,. 

Bugwood.org 

48.. John.H..Ghent,.USDA.Forest.Service,.Bugwood.org 

49.. Melody.Keena,.USDA.Forest.Service,.Bugwood.org 

50.. Michael.Bohne,.Bugwood.org 

51.. Kenneth.R..Law,.USDA.APHIS.PPQ,.Bugwood.org 

52.. Steven.Katovich,.USDA.Forest.Service,.Bugwood.org 

53.. Dennis.Haugen,.USDA.Forest.Service,.Bugwood.org 

54.. Dean.Morewood,.Health.Canada,.Bugwood.org 

55.. Natasha.Wright,.Florida.Department.of.Agriculture.and. 

Consumer.Services,.Bugwood.org 

56.. Donald.Duerr,.USDA.Forest.Service,.Bugwood.org 

57.. James.Solomon,.USDA.Forest.Service,.Bugwood.org 

58.. University.of.Arkansas.Forest.Entomology.Lab.Archive,. 

University.of.Arkansas,.Bugwood.org 

59.. Claude.Moffet,.Natural.Resources.Canada,.Canadian. 

Forest.Service,.Laurentian.Forestry.Centre 

60.. Larry.R..Barber,.USDA.Forest.Service,.Bugwood.org 

61.. USDA.Agricultural.Research.Service.Archive,.USDA. 

Agricultural.Research.Service,.Bugwood.org 

62.. Linda.Haugen,.USDA.Forest.Service,.Bugwood.org 

63.. USDA.Forest.Service.–.Region.8.–.Southern.Archive,. 

USDA.Forest.Service,.Bugwood.org 

64.. Natural.Resources.Canada,.Canadian.Forest.Service,. 

http://cfs.nrcan.gc.ca/ 

65.. Paul.H..Peacher,.USDA.Forest.Service,.Bugwood.org 

66.. David.J..Moorhead,.University.of.Georgia,.Bugwood.org 

67.. Andrej.Kunca,.National.Forest.Centre.–.Slovakia,. 

Bugwood.org 

68.. Louis-Michel.Nageleisen,.Département.de.la.Santé.des. 

forêts,.Bugwood.org 

69.. USDA.Forest.Service.–.Northeastern.Area.Archive,.USDA. 

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70.. André.Carpentier,.Natural.Resources.Canada,.Canadian. 

Forest.Service,.Laurentian.Forestry.Centre 

71.. USDA.Forest.Service.–.North.Central.Research.Station. 

Archive,.USDA.Forest.Service,.Bugwood.org 

72.. Tom.Coleman,.USDA.Forest.Service,.Bugwood.org 

73.. Manfred.Mielke,.USDA.Forest.Service,.Bugwood.org 

74.. Milan.Zubrik,.Forest.Research.Institute.–.Slovakia,. 

Bugwood.org 

75.. Jeffrey.Fengler,.Connecticut.Agricultural.Experiment. 

Station.Archive,.Connecticut.Agricultural.Experiment. 

Station,.Bugwood.org 

76.. Ferenc.Lakatos,.University.of.West-Hungary,.Bugwood. 

org 

77.. USDA.Forest.Service.Archive,.USDA.Forest.Service,. 

Bugwood.org 

78.. Haruta.Ovidiu,.University.of.Oradea,.Bugwood.org 

79.. Ronald.F..Billings,.Texas.A&M.Forest.Service,.Bugwood. 

org 

80.. Mark.Robinson,.USDA.Forest.Service,.Bugwood.org 

81.. Rusty.Haskell,.University.of.Florida,.Bugwood.org 

82.. William.A..Carothers,.USDA.Forest.Service,.Bugwood.org 

83.. Robert.L..Anderson,.USDA.Forest.Service,.Bugwood.org 

84.. Tamla.Blunt,.Colorado.State.University,.Bugwood.org 

85.. Petr.Kapitola,.State.Phytosanitary.Administration,. 

Bugwood.org 

86.. Minnesota.Department.of.Natural.Resources.Archive,. 

Minnesota.Department.of.Natural.Resources,.Bugwood. 

org 

87.. Steven.Valley,.Oregon.Department.of.Agriculture,. 

Bugwood.org 

88.. Fabio.Stergulc,.Università.di.Udine,.Bugwood.org 

89.. Bruce.Watt,.University.of.Maine,.Bugwood.org 

90.. Edward.L..Barnard,.Florida.Department.of.Agriculture. 

and.Consumer.Services,.Bugwood.org 

91.. Curtis.Utley,.CSUE,.Bugwood.org 

92.. J.R..Baker.&.S.B..Bambara,.North.Carolina.State. 

University,.Bugwood.org 

93.. Ned.Tisserat,.Colorado.State.University,.Bugwood.org 

94.. Roland.J..Stipes,.Virginia.Polytechnic.Institute.and.State. 

University,.Bugwood.org 

95.. Shari.Halik,.University.of.Vermont 

96.. Ronald.S..Kelley,.Vermont.Department.of.Forests,.Parks. 

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9.0 

AcronyMs 

CFIA. . Canadian.Food.Inspection.Agency 

CFS. . Canadian.Forest.Service 

DCH. . Department.of.Canadian.Heritage 

FGCA.. Forest.Gene.Conservation.Association 

FIAS. . Forest.Invasive.Alien.Species 

. 

ISC. . Invasive.Species.Centre 

ISRI. . Invasive.Species.Research.Institute 

NRCan. Natural.Resources.Canada 

GISD. . Global.Invasive.Species.Database 

MNDM. Ministry.of.Northern.Development.and.Mines 

MOE. . Ministry.of.the.Environment 

NCC. . Nature.Conservancy.of.Canada 

OFAH.. Ontario.Federation.of.Anglers.and.Hunters 

OMAFRA. Ontario.Ministry.of.Agriculture.and.Rural.Affairs 

OMNR. Ontario.Ministry.of.Natural.Resources 

OWA. . Ontario.Woodlot.Association 

PMRA.... Pest.Management.Regulatory.Agency 

USDA.. United.States.Department.of.Agriculture 

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