United States Department of Agriculture

Animal and Plant Health Inspection Service

Plant Protection and Quarantine

In order to prevent the introduction of quarantine pests into the United States, § 319.37-2a allows 

the APHIS Administrator to designate the importation of certain taxa of plants for planting as not 

authorized pending pest risk analysis (NAPPRA). APHIS has determined that the following 

plant taxa should be added to the NAPPRA category. In accordance with paragraph (b)(1) of that 

section, this data sheet details the scientific evidence APHIS evaluated in making the 

determination that the taxa are hosts of a quarantine pest.

Plants for Planting Quarantine Pest Evaluation Data Sheets

Table of Contents

Click on a quarantine pest name below to view the corresponding data sheet:

Anoplophora glabripennis Celtis

Cercidiphyllum

Koelreuteria

Tilia

Chrysanthemum stem Callistephus

necrosis virus (CSNV) Chrysanthemum (including syn. Dendranthema)

Eustoma

Dendroctonus micans Pseudotsuga

Moniliophthora perniciosa Arrabidaea

Bixa

Herrania

Theobroma

Monochamus alternatus Cedrus

Peanut clump virus and Arachis

Indian peanut clump virus Eleusine

Pennisetum
Setaria

Triticum

Zea

Phytophthora kernoviae Annona

Camellia

Drimys

Fagus

Gevuina

Hedera

Ilex

Leucothoe

Liriodendron

Lomatia

Magnolia (Michelia)

Pieris

Podocarpus

Quercus

Rhododendron

Sequoiadendron (Sequoia)

Vaccinium

Puccinia buxi Buxus
United States Department of Agriculture

Animal and Plant Health Inspection Service

Plant Protection and Quarantine

Plants for Planting Quarantine Pest Evaluation Data Sheet 

January 9th, 2013

In order to prevent the introduction of quarantine pests into the United States, § 319.37-2a allows 

the APHIS Administrator to designate the importation of certain taxa of plants for planting as not 

authorized pending pest risk analysis (NAPPRA). APHIS has determined that the following 

plant taxa should be added to the NAPPRA category. In accordance with paragraph (b)(1) of that 

section, this data sheet details the scientific evidence APHIS evaluated in making the 

determination that the taxa are hosts of a quarantine pest.

Quarantine Pest: Anoplophora glabripennis

Hosts: Acer spp., Aesculus spp., Albizia spp., Alnus spp., Betula spp., Carpinus

spp., Celtis spp., Cercidiphyllum spp., Elaeagnus spp., Fagus spp., Fraxinus

spp., Hibiscus spp., Koelreuteria spp., Liquidambar spp., Malus spp., Melia

spp., Morus spp., Platanus spp., Populus spp., Prunus spp., Pyrus spp.,

Robinia spp., Salix spp., Sophora spp., Sorbus spp., Tilia spp., Toona spp.,

Ulmus spp.

Status: This pest and most of these hosts were regulated under a Federal Order dated 

April 1, 2011. Cut flowers and greenery are also covered by the April 1, 2011 

Federal Order, but are not included in this action under NAPPRA.

__________________________________________________________________________

Taxonomy and description of the pest:

Anoplophora glabripennis Motschulsky (Coleoptera: Cerambycidae) 

Common name – Asian longhorned beetle (ALB) 

Known distribution:

A. glabripennis is native to China,1,3,6,8,9,10,13the Republic of Korea,1,3,6,8,10 and the 

Democratic People’s Republic of Korea1,3,6,10
It has been detected in Austria, France, Italy, Germany, 1,3,6,7

Switzerland5, the 

Netherlands4, and Canada1. Belgium has declared A. glabripennis as eradicated5. 

A.glabripennis has been detected in several U.S. states since 1996; however, current ALB 

infestations in New York, Massachusetts, New Jersey and Ohio are under official control 

and eradication.

Biology of the pest:

A. glabripennis generally takes one year to complete its life cycle, although two years is 

common as larvae must reach a critical weight before overwintering to induce pupation 

the next summer.6 Adult Asian longhorned beetles live for approximately a month3and 

have been observed from April to December, with peak activity occurring from May to 

July.6 Adult beetles feed on twigs, petioles, and veins of leaves for 10–15 days before 

initiating oviposition, typically on the upper trunk and main branches of host trees.6

Female beetles chew a funnel-shaped oviposition pit through the bark to oviposit a single 

egg, which will hatch after 1–2 weeks if laid during the summer months.6Females lay at 

least 32 eggs.5 Developing larvae initially create a feeding gallery in the cambial region 

and then an oval-shaped tunnel in the sapwood and heartwood of host trees.6Frass is 

expelled from the tunnels near the original oviposition site.6Pupation occurs in late 

spring or early summer at the end of the larval tunnel.6Exit holes in the bark of host trees 

from emerging adult beetles usually measure 10–15 mm in diameter6 and adults typically 

remain on the tree from which they emerged or fly short distances to nearby trees.5

Damage potential of pest:

Except on fruit-bearing trees, feeding activity of adult beetles on twigs and foliage is 

considered of minor importance as most damage is associated with larval tunneling in the 

cambial and wood regions of host trees.6 After years of attack, larval feeding can disrupt 

the vascular tissues, weaken structural integrity, and severely reduce the wood quality of 

hosts.6Symptoms of infested trees may include early abscission of leaves, thinning 

crowns, limb die-back, and breakage when subject to stress.13 Both healthy and stressed 

trees of varying sizes are candidates for attack by the Asian longhorned beetle and tree 

death is a common outcome of infestation.6

In addition to natural forests, 35% of urban trees in the United States are considered at 

risk for infestation by A. glabripennis.

4

If every urban locale in the conterminous United 

States were infected with the beetle, predicted consequences include a 34.9% loss of 

canopy cover, 30.3% total tree loss (1.2 billion trees), and $669 billion in lost value.9This 

estimate is likely to be low, as it does not include the threats to hardwood (lumber and 

maple sugar) and recreational industries.8

Control: 
Because larvae bore deep into the wood of host trees, biological or chemical pesticides 

are not effective methods of control.7 However, imidacloprid, a systemic neonicotinyl 

insecticide, injected into the soil or directly into host trees shows promise in controlling 

the beetle.7Imidacloprid causes significant mortality of adult beetles feeding on tree 

foliage due to ingestion and contact toxicity and of all exposed life stages feeding within 

the tree.10,13 Although its use does not provide complete control of A. glabripennis, it is 

useful as part of an integrated eradication or management program.10

Through movement of infested wood materials (pallets and crating from China, 

firewood), the potential for movement to other areas is high for this pest.7 Currently, the 

only proven effective method for limiting spread is an aggressive eradication program 

that involves regulatory activities, surveying, removing and properly disposing of all trees 

with signs of beetle infestation or that are deemed susceptible to future infestation.8,10,11

Since 1997, the USDA Animal and Plant Health Inspection Service’s program to fight 

this beetle has cost approximately $421 million. 

Known host range:

Acer spp.,1,2,12 Aesculus spp.,1,2,12 Albizia spp.,1,2,12 Alnus spp.,1,2,12 Betula spp.,1,2,12

Carpinus spp.,1,6 Celtis spp.,1,2,12 Cercidiphyllum spp.,1,2,12 Elaeagnus spp.,1,12 Fagus

spp.,

1 Fraxinus spp.,1,2,12 Hibiscus spp.,1,10 Koelreuteria spp.,1,12 Liquidambar spp.,1

Malus spp.,1,12 Melia spp.,1,12 Morus spp.,1,12 Platanus spp.,1,2,12 Populus spp.,1,2,12 Prunus

spp.,1,12 Pyrus spp.,1,12 Robinia spp.,1,12 Salix spp.,1,2,12 Sophora spp.,1,3 Sorbus spp.,1,2,12 

Tilia spp.,

1,12 Toona spp.,1 Ulmus spp.,1,2,12

Action under NAPPRA: 

The importation of the following plants for planting genera, excluding seeds that are hosts 

of A. glabripennis, is not authorized pending a pest risk analysis (NAPPRA) from all 

countries:

Koelreuteria.

The importation of the following plants for planting genera, excluding seeds that are hosts 

of A. glabripennis, is not authorized pending a pest risk analysis (NAPPRA) from all 

countries, except Canada:

Celtis, Tilia.

The importation of the following plants for planting genera, excluding seeds that are hosts 

of A. glabripennis, is not authorized pending a pest risk analysis (NAPPRA) from all 

countries, except Netherlands:

Cercidiphyllum.
The importation of the following plants for planting genera that are hosts of A. 

glabripennis is currently regulated under 7 CFR 319.37 and these genera are therefore 

not included, at this time, as potential candidates for listing on this NAPPRA data sheet:

Acer, Aesculus, Albizia, Alnus, Betula, Carpinus, Elaeagnus, Fagus, 

Fraxinus, Hibiscus, Liquidambar, Malus, Melia, Morus, Platanus, Populus, 

Prunus, Pyrus, Robinia, Salix, Sophora, Sorbus, Toona, and Ulmus.

References:

1. Canadian Food Inspection Agency. D-11-01. "Phytosanitary Requirements for Plants for 

Planting and Fresh Decorative." 1 April 2011. Canadian Food Inspection Agency. 4 

May 2012. 

http://members.wto.org/crnattachments/2011/sps/CAN/11_0373_00_e.pdf.

2. Code of Federal Regulations (CFR). 7§301.51-2(a). Domestic quarantine notices, Asian 

Longhorned Beetle, Regulated articles. Accessed date: May 8, 2012. 

http://ecfr.gpoaccess.gov/cgi/t/text/textidx?c=ecfr&sid=e7c42c7b63be222072ed48aa4d40e7ab&rgn=div8&view=text&node

=7:5.1.1.1.2.8.5.2&idno=7

3. EPPO. Data Sheets on Quarantine Pests: Anoplophora glabripennis. European and 

Mediterranean Plant Protection Organization. Accessed 16 December 2011. 

http://www.eppo.fr/QUARANTINE/insects/Anoplophora_glabripennis/ANOLGL_ds.

pdf

4. EPPO. (2010). EPPO Reporting Service. Paris: European and Mediterranean Plant 

Protection Organization. Published on 1 November 2010. Accessed on 8 May 2012 

at: http://archives.eppo.int/EPPOReporting/2010/Rse-1011.pdf

5. EPPO. (2011). EPPO Reporting Service. Paris: European and Mediterranean Plant 

Protection Organization. Published on 1 September 2011. Accessed on 8 May 2012 

at: http://archives.eppo.int/EPPOReporting/2011/Rse-1109.pdf

6. Haack, R.A., Herard, F., Sun, J., and Turgeon, J.J. 2010. Managing invasive populations 

of Asian longhorned beetle and citrus longhorned beetle: a worldwide perspective. 

Annual Review of Entomology. 55: 521-546. 

7. Herard, F., Ciampitti, M., Maspero, M., Krehan, H., Benker, U., Boegel, C., Schrage, R., 

Bouhot-Delduc, L., and P. Bialooki. 2006. Anoplophora species in Europe: 

infestations and management. Bulletin OEPP/EPPO Bulletin. 36: 470-474.

8. Keena, M.A. 2006. Effects of temperature on Anoplophora glabripennis (Coleoptera: 

Cerambycidae) adult survival, reproduction, and egg hatch. Environmental 

Entomology. 35 (4): 912-921.

9. Nowak, D.J., Pasek, J.E., Sequeira, R.A., Crane, D.E., and V.C. Mastro. 2001. Potential 

effect of Anoplophora glabripennis (Coleoptera: Cerambycidae) on urban trees in the 

United States. Journal of Economic Entomology. 94(1): 116-122.

10. Poland, T., Haack, R., Petrice, T., Miller, D., Bauer, L., and Gao, R. 2006. Field 
evaluations of systemic insectices for control of Anoplophora glabripennis

(Coleoptera: Cerambycidae) in China. Journal of Economic Entomology. 99(2): 383-

392.

11. Poland, T.M., Haack, R.A., and T.R. Petrice. 1998. Chicago joins New York in battle 

with the Asian longhorned beetle. Newsletter of the Michigan Entomological Society. 

43: 15-17.

12. Sawyer, A. 11 March 2011. Asian Longhorned Beetle: Annotated Host List. USDAAPHIS-PPQ, Center for Plant Health Science and Technology, Otis Laboratory. 

13. Wang, B., Gao, R., Mastro, V.C., and R.C. Reardon. Toxicity of four systemic 

neonicotinoids to adults of Anoplophora glabripennis (Coleoptera: Cerambycidae). 

Journal of Economic Entomology. 98(6): 2292-230
United States Department of Agriculture

Animal and Plant Health Inspection Service

Plant Protection and Quarantine

Plants for Planting Quarantine Pest Evaluation Data Sheet 

January 9th, 2013

In order to prevent the introduction of quarantine pests into the United States, § 319.37-2a allows 

the APHIS Administrator to designate the importation of certain taxa of plants for planting as not 

authorized pending pest risk analysis (NAPPRA). APHIS has determined that the following 

plant taxa should be added to the NAPPRA category. In accordance with paragraph (b)(1) of that 

section, this data sheet details the scientific evidence APHIS evaluated in making the 

determination that the taxa are hosts of a quarantine pest.

Quarantine Pest: Chrysanthemum stem necrosis virus (CSNV)

Hosts: Callistephus spp., Chrysanthemum spp. (including syn. Dendranthema spp.), 

Eustoma spp., Solanum spp. (including syn. = Lycopersicon spp.)

Status: Chrysanthemum spp. is regulated under 7 CFR 319.37 from all countries to 

prevent the introduction and establishment of the causal agent of Chrysanthemum 

white rust.

__________________________________________________________________________

Taxonomy and description of the pest:

Chrysanthemum stem necrosis virus (CSNV) is a single stranded RNA in the genus 

Tospovirus of the Bunyaviridae family.

Known distribution:

Brazil1, 5, Japan8

This pest is not known to occur in the United States.

Biology of the pest:

On chrysanthemum, CSNV may cause mild to severe necrotic streaks on the stem, wilting 

of leaves and stem, and chlorotic or necrotic spots and rings on leaves13. Peduncles and 

floral receptacles may also become necrotic5. Infection often leads to complete necrosis of 
the stem and wilting of sections of the plant13

. CSNV symptoms on chrysanthemum are 

similar to those of Tomato spotted wilt virus (TSWV); however, CSNV symptoms are 

generally more severe to the trained eye6.

CSNV is transmitted by thrips – insects of the family Thripidae (Thysanoptera). The 

widespread thrips species Frankliniella occidentalis and Frankliniella schultzei are 

vectors of CSNV1. F. occidentalis (common name: Western flower thrips) attacks a wide 

range of plants, including species susceptible to CSNV1, 3, 9. F. schultzei (common name: 

Common blossom thrips) is often found on plants in international trade14. Both F. 

occidentalis and F. schultzei are common greenhouse pests and are present in the United 

States3, 7. Seed transmission of CSNV is not known to occur.

Damage potential of pest:

CSNV was first recognized in Brazil, where losses due to the virus have not been 

quantified to date5, 6. CSNV was subsequently discovered in the Netherlands, England, 

Slovenia, and Japan2, 8, 10, 13. Concerns about potential economic losses and environmental 

impacts led to successful eradication campaigns in Europe (Netherlands, Slovenia, and 

United Kingdom)6, 10, 12, 13. The potential impact if CSNV became established in the 

United States is difficult to estimate but could be substantial.

Control:

Plants suspected of CSNV infection should be laboratory tested for reliable diagnosis4. 

Control measures for CSNV must be directed at exclusion of the thrips vectors. Control 

of thrips can be challenging and is best achieved through an integrated pest management 

strategy, which may include biological and chemical controls. Predatory insects and mites 

help to manage certain plant-feeding thrips species with varying degrees of success4. 

Unfortunately, chemical controls are minimally effective against thrips, and resistance is 

persistent3, 6. Proper cultural control and greenhouse hygiene may help to reduce thrips 

populations and mitigate the spread of CSNV.

Known host range:

Callistephus spp.

9

, Chrysanthemum spp. (including syn. Dendranthema spp.)6,10,13, 

Eustoma spp.

9

, Solanum spp. (including syn. = Lycopersicon spp.)1,6,9,11. 

Action under NAPPRA:

The importation of the following plants for planting genera, excluding seeds, which are 

hosts of CSNV, is not authorized pending pest risk analysis (NAPPRA) from all 

countries:

Callistephus, Eustoma.
The importation of the following plants for planting genera, excluding seeds, which are 

hosts of CSNV, are not authorized pending pest risk analysis (NAPPRA) from all 

countries, except Canada:

Chrysanthemum (including syn. Dendranthema).

The importation of the following plants for planting that are hosts of CSNV is currently 

regulated under 7 CFR 319.37 and this genus is therefore not included, at this time, as a 

potential candidate for listing on this NAPPRA data sheet:

Solanum (including syn. = Lycopersicon). 

References:

1. Bezerra, I. C., R. d. O. Resende, L. Pozzer, T. Nagata, R. Kormelink, and A. C. De Ávila. 

1999. Increase of tospoviral diversity in Brazil with the identification of two new 

tospovirus species, one from chrysanthemum and one from zucchini. 

Phytopathology. 89:823-830

2. Boben, J., N. Mehle, M. Pirc, I. Mavrič Pleško, and M. Ravnikar. 2007. New molecular 

diagnostic methods for detection of Chrysanthemum stem necrosis virus (CSNV). 

Acta Biologica Slovenica. 50:41-51

3. Cloyd, R. A. 2009. Western flower thrips (Frankliniella occidentalis) management on 

ornamental crops grwon in greenhouses: Have we reached an impasse? Pest Technol. 

3:1-9

4. Dreistadt, S. H., P. A. Phillips, and C. A. O'Donnell. 2007. Thrips. U Calif Agriculture

and Natural Resources Pest Notes. 7249:1-8

5. Duarte, L. M., E. B. Rivas, M. A. V. Alexandre, A. C. De Ávila, T. Nagata, and C. M. 

Chagas. 1995. Chrysanthemum stem necrosis caused by a possible novel tospovirus. 

J. Phytopathol. 143:569-571

6. EPPO. 2005. Data sheets on quarantine pests: Chrysanthemum stem necrosis tospovirus. 

EPPO Bulletin. 35:409-412

7. ITIS. Frankliniella schultzei (Trybom, 1910). Integrated Taxonomic Information System. 

Accessed December 16, 2011 at 

http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=

695334

8. Matsuura, S., K. Kubota, and M. Okuda. 2007. First report of Chrysanthemum stem 

necrosis virus on chrysanthemums in Japan. Plant Dis. 91:468

9. Momonoi, K., J. Moriwaki, and T. Morikawa. 2011. Stem necrosis of aster and Russell 

prairie gentian caused by Chrysanthemum stem necrosis virus. J. Gen. Plant Pathol. 

77:142-146

10. Mumford, R. A., B. Jarvis, J. Morris, and A. Blockley. 2003. First report of 

Chrysanthemum stem necrosis virus (CSNV) in the UK. Plant Pathol. 52:779
11. Nagata, T., R. d. O. Resende, E. W. Kitajima, H. Costa, A. K. Inoue-Nagata, and A. C. 

De Ávila. 1998. First report of natural occurrence of zucchini lethal chlorosis 

tospovirus on cucumber and chrysanthemum stem necrosis tospovirus on tomato in 

Brazil. Plant Dis. 82:1403

12. Ravnikar, M., N. Vozelj, I. Mavriè, S. D. Švigelj, M. Zupanèiè, and N. Petroviè. 2003. 

Detection of Chrysanthemum stem necrosis virus and Tomato spotted wilt virus in 

chrysanthemum. Abstracts 8th International Congress of Plant Pathology

13. Verhoeven, J. T. J., J. W. Roenhorst, I. Cortes, and D. Peters. 1996. Detection of a novel 

tospovirus in chrysanthemum. Acta Horticulturae. 432:44-53

14. Vierbergen, G., and W. P. Mantel. 1991. Contribution to the knowledge of Frankliniella 

schultzei (Thysanoptera: Thripidae). Entomologische Berichten (Amsterdam). 51:7-

12
United States Department of Agriculture

Animal and Plant Health Inspection Service

Plant Protection and Quarantine

Plants for Planting Quarantine Pest Evaluation Data Sheet 

January 9th, 2013

In order to prevent the introduction of quarantine pests into the United States, § 319.37-2a allows 

the APHIS Administrator to designate the importation of certain taxa of plants for planting as not 

authorized pending pest risk analysis (NAPPRA). APHIS has determined that the following 

plant taxa should be added to the NAPPRA category. In accordance with paragraph (b)(1) of that 

section, this data sheet details the scientific evidence APHIS evaluated in making the 

determination that the taxa are hosts of a quarantine pest.

Quarantine Pest: Dendroctonus micans Kugelann, 1794

Hosts: Abies spp., Larix spp., Picea spp., Pinus spp., Pseudotsuga spp.

__________________________________________________________________________

Taxonomy and description of the pest:

Dendroctonus micans (Coleoptera: Scolytidae)

Common name – Great spruce bark beetle 

Known distribution:

Australia11, Austria1, Belarus2, Belgium1, Bosnia and Herzegovina1,2, Bulgaria5,9, China2, 

7

, Croatia1,5, Czech Republic5, Denmark1,3, Estonia5,7, Finland2, France2,8, Georgia7, 

Germany2, Greece2,7, Hungary5, Ireland9, Italy11, Japan10, Latvia1,5, Lithuania1,2, 

Luxembourg1,7, Mongolia2,9, Montenegro9, Netherland1, Norway1,7, Poland5,11, Portugal9, 

Romania11, Russia2,9, Sakhalin2, Serbia and Montenegro1, Slovakia1,5, Slovenia5,11, 

Spain2,7, Sweden11, Switzerland11, Turkey3,9, Ukraine3, United Kingdom1,2

Biology of the pest:

D. micans has a robust, cylindrical, black or dark brown body, covered with tiny 

yellowish hair. The time required to complete one generation in field ranges from 1 to 3 

years, depending on the temperature of the environment7. The typical gender ratio is 1 

male/10 females but can be as high as 1 male/45 females1. Mating of the pest occurs 
under the bark. After mating, some adult females remain beneath the bark and simply 

initiate new galleries nearby2. Egg galleries are constructed primarily in the inner bark 

tissue and females lay between 100-150 eggs in a cluster7. Other females emerge and 

attach elsewhere on the same tree or fly to new host trees. Adult flight occurs through 

much of the summer when the threshold temperature of 20-23°C is met7. Mated females 

construct individual egg galleries in living trees from April to November, depending on 

local conditions2. 

Damage potential of pest:

D. micans normally occurs at low levels and causes little tree mortality. However, when 

several females oviposit close to each other and the individual galleries coalesce, this can 

cause extensive injury to the tree1. Outbreaks can result in widespread tree mortality as a 

result of the girdling action of larval feeding and oviposition1. This can take place over a 

period of several years until the whole tree is destroyed. As D. micans extended its range 

westward into Europe and southwestern Asia during the late 1900s, outbreaks occurred in 

more than 200,000 ha of spruce forest2. Normally, D. micans only colonizes green 

standing trees, but it will attack trees that are stressed as a result of logging damage, frost, 

snow, wind, lightning, poor soil nutrition and drought1. 

Known host range:

Abies spp.,

1,3,7,9 Larix spp.,1,3,7,9 Picea spp.,1,3,9 Pinus spp.,1,3,7,9 Pseudotsuga spp.1,3,7,9

Action under NAPPRA:

The importation of the following plants for planting genus, excluding seed a host of D. 

micans, is not authorized pending pest risk analysis (NAPPRA) from all countries, 

except Canada and India:

Pseudotsuga.

The importation of the following plants for planting genera that are hosts of D. micans is 

currently regulated under 7 CFR 319.37 and these genera are therefore not included, at 

this time, as potential candidates for listing on this NAPPRA data sheet:

Abies, Larix, Picea, and Pinus.

References:

1. CABI (2006). Crop Protection Compendium (2006 ed.) [CD]. Wallingford, UK: CAB 

International. Access Date: 28 July 2011. Current online version at 

http://www.cabi.org/cpc/
2. EcoPort: Dendroctonus micans. (n.d.). EcoPort Foundation. Access Date: 28 July 2011.

http://ecoport.org/ep?Arthropod=26421&entityType=AR****&entityDisplayCategor

y=full

3. EPPO (n.d.). Data Sheets on Quarantine Pests: Dendroctonus micans. European and 

Mediterranean Plant Protection Organization. Access Date: 28 July 2011. 

http://www.eppo.org/QUARANTINE/insects/Dendroctonus_micans/DENCMI_ds.p

df

4. European Nature Information System (EUNIS): Dendroctonus micans. (n.d.). 

Copenhagen, Denmark: European Environment Agency. Access Date: 28 July 2011. 

http://eunis.eea.europa.eu/speciesfactsheet.jsp?idSpecies=268554&idSpeciesLink=268554

5. Fauna Europaea: Dendroctonus micans. (2007, April 19). Access Date: 28 July 2011. 

http://www.faunaeur.org/full_results.php?id=250017

6. Great Britain Forestry Commission (2002, May). Dendroctonus micans - a guide for 

Forest Managers on control techniques. Plant Health Leaflet, 9. Access Date: 28 July 

2011. http://www.forestry.gov.uk/pdf/ph9.pdf/$FILE/ph9.pdf

7. Haack, R. A. (2001, February 14). Exotic Forest Pest Information System for North 

America: Dendroctonus micans. North American Forest Commission. Access Date: 

28 July 2011. 

http://spfnic.fs.fed.us/exfor/data/pestreports.cfm?pestidval=35&langdisplay=english

8. The Bugwood Network: Dendroctonus micans. (2006, January 17). University of 

Georgia. Access Date: 28 July 2011. 

http://www.forestryimages.org/search/action.cfm?q=Dendroctonus%20micans&Start

=1&results=30

9. USDA-APHIS-PPQ-CPHST, CAPS. Venette, R. C. (Ed.) (2008, June (Updated 2010, 

July)). Pine Commodity-based Survey Reference. 1-221. United States Department 

of Agriculture, Animal and Plant Health Inspection Service, Plant Protection and 

Quarantine, Center for Plant Health Science and Technology, Cooperative 

Agriculture Pest Survey. Access Date: 28 July 2011. 

http://caps.ceris.purdue.edu/survey/manual/pine_reference

10. Walker, K. (2007). Pests and Diseases Image Library (PaDIL): Dendroctonus micans.

Access Date: 28 July 2011. 

http://www.padil.gov.au/viewPestDiagnosticImages.aspx?id=467

11. Zipcode Zoo: Dendroctonus micans. (2007, December 28). BayScience Foundation. 

Access Date: 28 July 2011. 

http://zipcodezoo.com/Animals/D/Dendroctonus_micans/Default.asp
 United States Department of Agriculture

 Animal and Plant Health Inspection Service

 Plant Protection and Quarantine

Plants for Planting Quarantine Pest Evaluation Data Sheet 

January 9th, 2013

In order to prevent the introduction of quarantine pests into the United States, § 319.37-2a allows 

the APHIS Administrator to designate the importation of certain taxa of plants for planting as not 

authorized pending pest risk analysis (NAPPRA). APHIS has determined that the following 

plant taxa should be added to the NAPPRA category. In accordance with paragraph (b)(1) of that 

section, this data sheet details the scientific evidence APHIS evaluated in making the 

determination that the taxa are hosts of a quarantine pest.

Quarantine Pest: Moniliophthora perniciosa

Hosts: Arrabidaea spp., Bixa spp., Capsicum spp., Herrania spp., Solanum spp., 

Theobroma spp.

__________________________________________________________________________

Taxonomy and description of the pest:

Moniliophthora perniciosa is a basidiomycete fungal pathogen in the Marasmiaceae 

family. The fungus is highly pathogenic to Theobroma cacao and is commonly referred to 

as the “witches’ broom fungus” due to the unusual appearance of shoots projecting from 

infected plants. M. perniciosa was first named Marasmius perniciosus20 and has also been 

called Crinipellis perniciosa1. Common names for witches’ broom disease of cacao 

include “balai de sorcihre du cacaoyer” (French), “cacao witches’ broom disease,” 

“Crinipellis pod rot,”13 “escoba de bruja del cacao” (Spanish), “hexenbesenkrankheit 

kakao” (German), “lagaratao,”14 and “vassoura de bruxa” (Portuguese).

Known distribution:

Bolivia11, 16, Brazil11, 16, Colombia11, 16, Ecuador11, 16, Grenada11, 16, Guyana11, 16, 

Panama16, Peru11, 16, Saint Lucia12, 16, Saint Vincent16 and Grenadines12, Suriname16, 20, 

Trinidad3, 16 and Tobago11, Venezuela11, 16.

This pest is not known to occur in the United States.
Biology of the pest:

The M. perniciosa disease cycle consists of two phases. In the biotrophic phase, the 

pathogen invades young, growing tissue causing hypertrophy (enlargement) and 

hyperplasia (proliferation) of the plant’s cells11. Symptoms of biotrophic infection include 

loss of apical dominance, proliferation of auxiliary shoots, and the formation of abnormal 

stems resulting in broom-like structures14. Infection of the cauliform flowers results in the 

formation of dry brooms and small parthenocarpic (seedless) fruits14. Under high 

humidity7, M. perniciosa basidiospores may infect any meristematic cacao tissues, 

including shoots, flowers, and developing fruits6. The basidiospores produce germ tubes, 

which allow direct penetration of leaves or entry through stomatal openings or the bases 

of damaged trichomes7, 19. Infected pods may display black or brown lesions, ooze, and 

deformities such as scabbing, pitting, abnormal shape, and malformed skin1. Additionally, 

leaves may show necrosis and seeds may fuse together and display lesions14. Necrosis and 

death of the infected tissues eventually lead to the formation of dry broom structures, 

which remain attached to the plant6.

During the saprotrophic phase of its disease cycle, M. perniciosa proliferates and 

colonizes necrotic or dead host tissue. Following alternating wet and dry periods2, small, 

pink basidiocarps (mushrooms) may form on any infected necrotic tissue and produce 

basidiospores, thus completing the disease cycle14. From a single dry broom, the 

formation of basidiocarps and production of basidiospores may occur repeatedly over 

many years

14

.

Damage Potential of Pest:

Witches’ broom is one of the two most devastating diseases of cacao in tropical 

America1. M. perniciosa is responsible for the failure of the cocoa industry in Suriname,

where it arrived in 189518, 20. The pathogen subsequently spread through the northern 

parts of South America, where it catastrophically affected cocoa production with yield 

reductions of 50-90%14. M. perniciosa arrived in Bahia, Brazil in 1989 and caused the 

country to fall from the world’s third largest producer of cocoa to a net importer14. 

In the United States, T. cacao is grown in Puerto Rico, Florida, and Hawaii. While there 

are currently relatively few T. cacao plantations in the U. S., significant efforts are 

underway to boost the cocoa industry in Hawaii. Analysts project that Hawaiian cacao 

farming could one day bring $2 to $34 million annually into the U. S. economy, making it 

the third-largest Hawaiian crop8. M. perniciosa must be prevented from entering the 

United States, especially Hawaii, where it could have serious consequences for the 

budding cacao industry. 

Control:

Control of M. perniciosa in cacao is a formidable task. Four general control strategies –

phytosanitation, chemical control, genetic resistance, and biological control – have been 
utilized with varying success. Phytosanitation (the removal and destruction of diseased 

plant parts) has remained the basis of witches’ broom control since the beginning of the 

twentieth century. Unfortunately, the strategy is tedious and expensive, and it may require 

up to 95% removal of infected tissue to achieve only 50% reduction in pod loss15.

Chemical control is not widely practiced in cacao production due to the associated high 

costs and environmental risks14. A number of cacao accessions carry genetic resistance to 

M. perniciosa, including two Peruvian clones that have been used extensively in breeding 

programs14. However, while these clones showed resistance in Brazil and Trinidad, they 

were susceptible in Ecuador, indicating that geographical variations within the pathogen 

impact resistance4. T. cacao endophytes are being studied for their potential utility as 

agents of biocontrol against M. perniciosa. For example, commercial formulations of 

Trichoderma stromaticum, a newly discovered T. cacao endophyte that parasitizes the 

saprophytic mycelia and basidiocarps of M. perniciosa, have been used in Brazil with 

some success14, 17. Endophytes represent a promising strategy for the control of witches’ 

broom, but more work must be done to identify the optimal environmental conditions for 

their colonization and suppression of M. perniciosa in nature14. 

Known host range:

Arrabidaea spp.10, Bixa spp.9, Capsicum spp.5, Herrania spp.3, 21, Solanum spp.5, 

Theobroma spp.1, 21.

Action under NAPPRA:

The importation of the following plants for planting genera, excluding seed but including 

cut flowers and greenery, hosts of M. perniciosa, is not authorized pending pest risk 

analysis (NAPPRA) from all countries:

Arrabidaea, Bixa, Herrania, Theobroma.

The importation of the following plants for planting genera, excluding seed but including 

cut flowers and greenery, that are hosts of M. perniciosa is currently regulated under 7 

CFR 319.37 and these genera are therefore not included, at this time, as potential 

candidates for listing on this NAPPRA data sheet:

Capsicum, Solanum.

References:

1. Aime, M. C., and W. Phillips-Mora. 2005. The causal agents of witches' broom and frosty 

pod rot of cacao (chocolate, Theobroma cacao) form a new lineage of 

Marasmiaceae. Mycologia. 97:1012-1022

2. Almeida, O. C., F. P. B. Chiacchio, and H. M. Rocha. 1997. Sobrevivência de Crinipellis 

perniciosa (Stahel) Singer em vassouras secas de cacaueiros (Theobroma cacao L.) 

do estado da Bahia. Agrotrópico. 9:23-28
3. Baker, R. E. D., and H. P. 1957. Witches' broom disease of cocoa (Marasmius 

perniciosus Stahel), Phytopathological Papers, vol. 2. Commonwealth Mycological 

Institute, Kew, U.K.

4. Bartley, B. G. D. 2001. The origin and compatibility relationships of the Scavina variety 

of Theobroma cacao L. Ingenic Newsletter. 6:23-24

5. Bastos, C. N., and H. C. Evans. 1985. A new pathotype of Crinipellis perniciosa (witches' 

broom disease) on solanaceous hosts. Plant Pathol. 34:306-312

6. Evans, H. C. 1980. Pleomorphism in Crinipellis perniciosa, causal agent of witches' 

broom disease of cocoa. Trans. Brit. Mycol. Soc. 74:515-526

7. Frias, G. A., L. H. Purdy, and R. A. Schmidt. 1991. Infection biology of Crinipellis 

perniciosa on vegetative flushes of cacao. Plant Dis. 75:552-556

8. Gomes, A. Hoping for a sweet harvest. The Honolulu Advertiser. January 25, 2010

9. Griffith, G. W., and J. N. Hedger. 1994. The breeding biology of biotypes of the witches' 

broom pathogen of cocoa, Crinipellis perniciosa. Heredity. 72:278-289

10. Griffith, G. W., and J. N. Hedger. 1994. Spatial distribution of mycelia of the liana (L-) 

biotype of the agaric Crinipellis perniciosa (Stahel) Singer in tropical forest. New 

Phytol. 127:243-259

11. Holliday, P. 1980. Fungus diseases of tropical crops. Cambridge University Press, 

Cambridge, 612 pp.

12. Kelly, P. L., R. Reeder, S. Rhodes, and N. Edwards. 2009. First confirmed report of 

witches' broom caused by Moniliophthora perniciosa on cacao, Theobroma cacao, in 

Saint Lucia. Plant Pathol. 58:798

13. Maddison, A. C., G. Macias, C. Moreira, R. Arias, and R. Neira. 1995. Cocoa production 

in Ecuador in relation to dry-season escape from pod rot caused by Crinipellis 

perniciosa and Moniliophthora roreri. Plant Pathol. 44:982-998

14. Meinhardt, L. W., J. Rincones, B. A. Bailey, M. C. Aime, G. W. Griffith, D. Zhang, and 

G. A. G. Pereira. 2008. Moniliophthora perniciosa, the causal agent of witches' 

broom disease of cacao: what's new from this old foe? Mol. Plant Pathol. 9:577-588

15. Rudgard, S. A., and D. R. Butler. 1987. Witches' broom disease in Rondonia, Brazil: pod 

infection in relation to pod susceptibility, wetness, inoculum, and phytosanitiation. 

Plant Pathol. 36:515-522

16. Rudgard, S. A., A. C. Maddison, and T. Andebrhan (ed.). 1993. Disease management in 

cocoa: comparative epidemiology of witches' broom. Chapman & Hall, London.

17. Samuels, G. J., R. Pardo-Schultheiss, P. K. Hebbar, R. D. Lumsden, C. N. Bastos, J. C. 

Costa, and J. L. Bezerra. 2000. Trichoderma stromaticum, sp. nov., a parasite of 

cacao witches' broom pathogen. Mycol Res. 104:760-764

18. Singer, R. 1942. A monographic study of the genera Crinipellis and Chaetocalathus. 

Lilloa. 8:441-534

19. Sreenivasan, T. N., and S. S. Dabydeen. 1989. Modes of penetration of young cocoa 

leaves by Crinipellis perniciosa. Plant Dis. 73:478-481

20. Stahel, G. 1915. Marasmius perniciosus nov. spec. Bulletin Departement van den 

Landbouw in Suriname. 33:1-26

21. Wood, G. A. R., and R. A. Lass. Cocoa, Fourth ed. Blackwell Science Ltd., Oxford, 620 

pp.
United States Department of Agriculture

Animal and Plant Health Inspection Service

Plant Protection and Quarantine

Plants for Planting Quarantine Pest Evaluation Data Sheet 

January 9th, 2013

In order to prevent the introduction of quarantine pests into the United States, § 319.37-2a allows 

the APHIS Administrator to designate the importation of certain taxa of plants for planting as not 

authorized pending pest risk analysis (NAPPRA). APHIS has determined that the following 

plant taxa should be added to the NAPPRA category. In accordance with paragraph (b)(1) of that 

section, this data sheet details the scientific evidence APHIS evaluated in making the 

determination that the taxa are hosts of a quarantine pest.

Quarantine Pest: Monochamus alternatus Hope, 1843

Hosts: Abies spp., Acer spp., Cedrus spp., Cryptomeria spp., Fagus spp., Liquidambar 

spp., Malus spp., Picea spp., Pinus spp., Quercus spp.

__________________________________________________________________________

Taxonomy and description of the pest:

Monochamus alternatus Hope (Coleoptera: Cerambycidae)

Synonym: Monochammus tesserula

Common names: Japanese pine sawyer, pine sawyer beetle, rusty pine longhorn

Known distribution:

China 7, 14, Hong Kong2, 7, Japan7, Republic of Korea2, 11, Laos7, 8, Taiwan Island2, 3, Viet 

Nam3, 7.

Biology of the pest:

Monochamus alternatus generally completes one generation per year4. Larvae overwinter 

within the tunnel, and begin to pupate in late March. Adults emerge in mid-April, but stay 

in host trees for 6-8 days. Peak of adult activity occurs in May. Adults prefer to feed on 

younger trees. Although most adults will not fly more than a few hundred meters they can 
disperse several kilometers9

. Populations of M. alternatus can spread up to 20 km/yr 

depending on the change in distribution of trees infested9. 

Weakened or newly felled trees are preferred for oviposition5. The female beetle lays 

white, sickle shaped eggs under the bark and the young round-headed borer larvae feed in 

the softer inner bark tissues4, 10. The larvae are white and opaque legless grubs averaging 

43mm in length4. After developing strong mandibles, the larvae start boring into the 

trunk. They continue to feed on trunk until maturity10. The mature larvae bore back to the 

sapwood area in the outer region of the trunk where they excavate a large chamber in 

which they pupate10. It is during the transformation to the adult stage when the pinewood 

nematode, which has been feeding and multiplying in the tree tissues, infests the tracheal 

system of the adult10. Adult maturation feeding consists of removal of the bark from 

shoots of host trees. The shoots eventually die leaving dead tips on the trees4. 

Damage potential of pest:

Monochamus alternatus is known to be one of the most destructive pests associated with 

pines in Japan. Even though, M. alternates is primarily known as a pest of pine, it will 

feed on a number of other plants in the Pinaceae and Cupressaceae families4. Adults 

cause damage to hosts by feeding on shoots and creating ovipositor sites. The larvae 

disrupt phloem as they feed10. M. alternatus is a vector of pinewood nematode, 

Bursaphelenchus xylophilus, in China and Japan and is known to be capable of carrying 

an average of 18,000 individual nematodes4. The interaction between M. alternates and 

B. xylophilus causes heavy loss of timber and international trade2. 

Known host range:

Abies spp.

3, 6

, Acer spp.

12

, Cedrus spp.

13

, Cryptomeria spp.

5

, Fagus spp.

5

, Liquidambar 

spp.

12

, Malus spp.

12

, Picea spp.

3,6

, Pinus spp.

5

, Quercus spp.

2

Action under NAPPRA:

The importation of the following plants for planting genera, excluding seed that are hosts 

of Monochamus alternates, is not authorized pending pest risk analysis (NAPPRA) from 

all countries, except Canada:

Cedrus.

The importation of the following plants for planting genera that are hosts of Monochamus 

alternates is currently regulated under 7 CFR 319.37 and these genera are therefore not 

included, at this time, as potential candidates for listing on this NAPPRA data sheet:

Abies, Acer, Cryptomeria, Fagus, Liquidambar, Malus, Picea, Pinus, 
Quercus.

References:

1. Bark Beetles: Monochamus alternatus. (n.d.). Cooperative Agriculture Pest Survey 

Program. Retrieved August 1st, 2011, from 

http://www.ceris.purdue.edu/napis/pests/barkb/monocfs.html

2. CABI. 2005. Crop Protection Compendium. CAB International. Accessed online 1st

August 2011 from

http://www.cabi.org/cpc/?compid=1&dsid=34719&loadmodule=datasheet&page=86

8&site=161.

3. Chen YM, Chao JT, 1998. Three species of long-horned beetles feeding on Taiwan red 

pine. Taiwan Journal of Forest Science, 13:373-376.

4. Ciesla, W. M. (2001, July 12). Exotic Forest Pest Information System for North America: 

Monochamus alternatus. North American Forest Commission. Retrieved February 

13, 2007, from 

http://spfnic.fs.fed.us/exfor/data/pestreports.cfm?pestidval=77&langdisplay=english

5. EcoPort: Monochamus alternatus. (n.d.). EcoPort Foundation. Retrieved February 13, 

2007, from http://www.ecoport.org

6. EPPO (n.d.). Data Sheets on Quarantine Pests: Bursaphelenchus xylophilus. 1-12. 

European and Mediterranean Plant Protection Organization. Retrieved February 13, 

2007, from 

http://www.eppo.org/QUARANTINE/nematodes/Bursaphelenchus_xylophilus/BUR

SXY_ds.pdf

7. EPPO Global Database, 2012. Eurpean and Mediterranean Plant Protection organization. 

Accessed December 27th, 2012. Available from:

http://gd3.eppo.int/organism.php/MONCAL

8. Gressitt JT, Rondon JA, von Breuning S, 1970. Cerambycid Beetles of Laos. Pacific 

Insects Monograph, 24:i-vi.

9. Kishi, Y. 1995. The pine wood nematode and the Japanese pine sawyer. Thomas 

Company Limited, Tokyo.

10. McLean, J. A. (1996, August 12). Pinewood Nematode in Jiangsu Province, People's 

Republic of China. University of British Columbia Forestry. Retrieved February 13, 

2007, from http://www.forestry.ubc.ca/fetch21/chinapwn/china96.html

11. Park NC, Moon YS, Lee SM, Yi CK, 1992. Distribution and oviposition activities of 

Monochamus alternatus Hope (Coleoptera: Cerambycidae) in Korea. Research 

Reports of the Forestry Research Institute (Seoul), No.44:142-150; [With English 

figures and tables]; 16 ref.

12. USDA-APHIS-PPQ-CPHST, CAPS. Venette, R. C. (Ed.) (2008, June (Updated 2010, 

July)). Pine Commodity-based Survey Reference. 1-221. United States Department 

of Agriculture, Animal and Plant Health Inspection Service, Plant Protection and 

Quarantine, Center for Plant Health Science and Technology, Cooperative 
Agriculture Pest Survey. Retrieved May 2, 2012, from 

http://caps.ceris.purdue.edu/survey/manual/pine_reference

13. Walker, K. (2006). Pests and Diseases Image Library (PaDIL): Pine Sawyer beetle.

Retrieved May 7, 2012, from http://www.padil.gov.au/pests-anddiseases/Pest/Main/135562

14. Wang SF, 1983. Monochamus alternatus Hope. In: Chinese Academy of Forestry, ed. 

Forest Insects in China. Beijing, China: Chinese Forestry Press, 284-285.
United States Department of Agriculture

Animal and Plant Health Inspection Service

Plant Protection and Quarantine

Plants for Planting Quarantine Pest Evaluation Data Sheet 

January 9th, 2013

In order to prevent the introduction of quarantine pests into the United States, § 319.37-2a allows 

the APHIS Administrator to designate the importation of certain taxa of plants for planting as not 

authorized pending pest risk analysis (NAPPRA). APHIS has determined that the following 

plant taxa should be added to the NAPPRA category. In accordance with paragraph (b)(1) of that 

section, this data sheet details the scientific evidence APHIS evaluated in making the 

determination that the taxa are hosts of a quarantine pest.

Quarantine Pests: Peanut clump virus 

Indian peanut clump virus

Hosts: Peanut clump virus: Arachis spp., Pennisetum spp., Saccharum spp.,

Sorghum spp., Triticum spp.

Indian peanut clump virus: Arachis spp., Eleusine spp., Triticum spp., 

Hordeum spp., Pennisetum spp., Setaria spp., Sorghum spp., Zea spp.

Status: Arachis spp. seeds are currently prohibited from Burkina Faso, Cote d’lvoire, 

Senegal and India (7 CFR 319.37-2). All other plant parts from the genus Arachis 

are prohibited from all countries excluding Canada under the Fabaceae 

prohibition in 7 CFR 319.37-2.

__________________________________________________________________________

Taxonomy and description of the pest:

Peanut clump virus (PCV) and Indian peanut clump virus (IPCV) are single stranded 

RNA viruses in the genus Pecluvirus of the family Virgaviridae[1].

Known distribution:

Peanut clump virus: Africa, Burkina Faso[2-4], Chad[5], Congo[5], Gabon[5], Côte 

d’Ivoire[3], Mali[5], Niger[3], Senegal[2, 6], Sudan[5]
Indian peanut clump virus: India[7], Pakistan[8]

These viruses are not known to occur in the United States.

Biology of the pest:

IPCV and PCV are thought to be cereal viruses that have coevolved as opportunistic 

pathogens of peanut when that crop was introduced to Africa and the Indian subcontinent. 

The viruses cause a “clump” disease in peanuts characterized by dark green-colored, 

miniaturized leaves, short internodes and dwarfing,[3, 5, 6]. IPCV causes a similar disease 

in peanut on the Indian subcontinent. Though the viruses cause similar diseases and share 

other similarities they are serologically and genomically distinct [9-11]. Both viruses may 

be transmitted through seed, soil and/or vegetative propagation [5]. 

In the soil, the viruses are transmitted by the obligate root endoparasite Polymyxa 

graminis

[12, 13]

. P. graminis is a plasmodiophorid protist responsible for the transmission 

of several very important plant viruses [12-15]. P. graminis heavily infests the roots of 

pearl millet, sorghum and sugarcane[5, 15]. These plant species are well-adapted to the P. 

graminis vector, and they support PCV multiplication[5]. Therefore, cultivation of pearl 

millet, sorghum and sugarcane on PCV-infested soils increases the virus inoculum. 

Resting P. graminis spores remain in the soil in the absence of plants, allowing PCV to 

persist between plantings. PCV-infested P. graminis spores may be dispersed by soil 

tillage and irrigation practices[5].

PCV and IPCV are regarded as high-risk pathogens for germplasm exchange because they 

are seed-transmitted at a high frequency in peanut[16, 17] and at lower rates in cereal and 

grain crops[5, 16]. In addition, low levels of viral inoculum may be maintained in seed 

stocks for several seasons[5]. PCV can also be propagated vegetatively through sugarcane 

cuttings or the rhizomes of grass weeds[5].

Damage potential of pest:

PCV is widely distributed in western Africa and is economically important in Niger, 

Burkina Faso, and Senegal[18]. Losses due to PCV can be significant, and the persistence 

of inoculum in the soil can lead to the abandonment of groundnut as a crop[19]. Similarly, 

IPCV has been reported to be economically important in groundnut grown on the Indian 

subcontinent[20]. In 1999, annual losses caused by clump disease in peanut globally were

estimated to exceed US$ 38 million[21]. Altogether, viral diseases on groundnut, chickpea, 

pigeonpea, sorghum and pearl millet in the semi-arid tropics result in yield losses worth 

over $700 million per year[22]. Because of its widespread occurrence, endemic nature and 

debilitating impact on crop yield, PCV and IPCV are a main focus of research in the 

semi-arid tropics[22].
Control:

Avoidance of peanut clump disease is best achieved through the use of clean seed. 

Likewise, groundnut seeds and sugarcane cuttings should be produced in disease-free 

areas. The pest may also be controlled by soil fumigation with nematicides that have 

fungicidal action[2]. 

The use of machinery for soil preparation and irrigation increases the impact and the 

spread of P. graminis and peanut clump disease. Therefore, machinery use should be 

restricted in infested fields[5]. Pearl millet seeds should be collected from the lower third 

of inflorescence, and collecting groundnut seeds from PCV-infected plots should be 

avoided[5]. Alternative cropping methods, such as trap cropping or crop rotations using 

non-hosts of Polymyxa (e.g., sunflower or mustard) may limit the spread of the disease[5].

Known host range:

Peanut clump virus: Arachis hypogaea[4, 12, 17], Pennisetum glaucum[5], , Saccharum 

officinarum[23], Sorghum arundinaceum[12, 24], Triticum aestivum[12]

Indian peanut clump virus: Arachis hypogaea[11, 20], Eleusine coracana[16],Triticum 

aestivum[25], Hordeum vulgare[25], Pennisetum glaucum[5], , Setaria italica[16], Sorghum 

bicolor[16], Zea mays[26]

Bold type indicates host with known seed transmission.

Action under NAPPRA:

The importation of seeds of the following plants for planting genera that are hosts of 

Peanut clump virus and/or Indian peanut clump virus, is not authorized pending a pest 

risk analysis (NAPPRA) from all countries:

Arachis, Eleusine.

The importation of seeds of the following plants for planting genera that are hosts of 

Peanut clump virus and/or Indian peanut clump virus, is not authorized pending a pest 

risk analysis (NAPPRA) from all countries, except those listed after the plant genus:

Pennisetum Australia, Germany

Setaria Netherland

Triticum Argentina, Canada, France, Germany, Mexico, New 

Zealand, United Kingdom, 
Zea Argentina, Austria, Brazil, Canada, Chile, Croatia, 

France, Germany, Guyana, Hungary, Italy, Mexico, 

Netherlands, New Zealand, Peru, Romania, Serbia, 

Spain, Turkey, United Kingdom

Peanut clump virus and/or Indian peanut clump virus are not known to be seed 

transmitted in the following taxa, and therefore the seed of these taxa are not included, at 

this time, as potential candidates for listing on this NAPPRA data sheet, but are currently 

regulated under 7 CFR 319.37:

Hordeum, Saccharum, and Sorghum.

The importation of vegetative parts of the following plants for planting genera that are 

hosts of Peanut clump virus and/or Indian peanut clump virus, is not authorized pending 

a pest risk analysis (NAPPRA) from all countries

Arachis, Eleusine, Pennisetum, Setaria.

The importation of vegetative parts of the following plants for planting genera that are 

hosts of Peanut clump virus and/or Indian peanut clump, is currently regulated under 7 

CFR 319.37 and vegetative parts of these genera are therefore not included, at this time, 

as potential candidates for listing on this NAPPRA data sheet:

Triticum, Zea, Saccharum, Sorghum, Hordeum.

References:

1. ICTV. International Congress of Virus Taxonomy. 2009 [cited 2012 May 29]; Available 

from: http://ictvonline.org/virusTaxonomy.asp?version=2009&bhcp=1.

2. Germani, G., J.-C. Thouvenel, and M. Dhery, Le rabougrissement de l'arachide: une 

maladie à virus au Sénégal et en Haute-Volta. Oléagineux, 1975. 30(6): p. 259-266.

3. Manohar, S.K., Dollet, M., Dubern, J., Gargani, D, Studies on variability of peanut clump 

virus: symptomatology and serology. J. Phytopathol., 1995. 143(4): p. 233-238.

4. Thouvenel, J.-C., M. Dollet, and C. Fauquet, Some properties of peanut clump, a newly 

discovered virus. Ann. Appl. Biol., 1976. 84(3): p. 311-320.

5. Dieryck, B., et al., Seed, soil and vegetative transmission contribute to the spread of 

pecluviruses in Western Africa and the Indian sub-continent. Virus Res., 2009. 

141(2): p. 184-189.

6. Bouhot, D., Observations sur quelques affections des plantes cultivées au Sénégal.

Agronomie Tropicale, 1967. 22: p. 888-890.

7. Reddy, D.V.R., R. Rajeshwari, N. Iisuka, D.E. Lesemann, B. L. Nolt and T. Goto, The 

occurrence of Indian peanut clump, a soil-borne virus disease of groundnuts 

(Arachis hypogaea) in India. Ann. Appl Biol., 1983. 102: p. 305-310.
8. Delfosse, P., M. Bashir, S. N. Malik and A.S. Reddy, Survey of groundnut virus diseases 

in Pakistan. International Arachis Newsletter, 1995. 15: p. 51-52.

9. ICTV. Index of Viruses - Pecluvirus In: ICTVdB - The Universal Virus Database, 

version 4. 2006 [cited 2012 May 29]; Available from: 

,http://www.ncbi.nlm.nih.gov/ICTVdb/Ictv/fs_index.htm.

10. Naidu, R.A., J. S.Miller, M. A. Mayo and A. S. Reddy, The nucleotide sequence of Indian 

peanut clump virus RNA 2. in Third Symp. Int. Working Group on Plant Viruses with 

Fungal Vectors. 1996. Dundee, Scotland.

11. Nolt, B.L., R. Rajeshwar, D.V.R. Reddy, N. Bharathan and S.K. Manohar, Indian peanut 

clump virus isolates: Host range,symptomatology, serological relationships,and 

some physical properties. Phytopathology, 1988. 78: p. 310-313.

12. Thouvenel, J.-C. and C. Fauquet, Further properties of peanut clump virus and studies on 

its natural transmission. Ann. Appl. Biol., 1981. 97: p. 99-107.

13. Ratna, A.S., A.S. Rao, A.S. Reddy, B. L. Nolt, D.V.R. Reddy, M. Vijayalakshmi and D. 

McDonald, Studies on transmission of Indian peanut clump virus disease by 

Polymyxa graminis. Ann. Appl.Biol., 1991. 118: p. 71-78.

14. Kanyuka, K., E. Ward, and M.J. Adams, Polymyxa graminis and the cereal viruses it 

transmits: a research challenge. Mol. Plant Pathol., 2003. 4(5): p. 393-406.

15. Legrève, A., Vanpee, B., Delfosse, P. and Maraite, H., Host range of tropical and subtropical isolates of Polymyxa graminis. Eur. J. Plant Pathol., 2000. 106(4): p. 379-

389.

16. Reddy, A.S., Hobbs, H. A., Delfosse, P., Murthy, A. K. and Reddy, D. V. R., Seed 

transmission of Indian peanut clump virus (IPCV) in peanut and millets. Plant Dis., 

1998. 82(3): p. 343-346.

17. Thouvenel, J.-C., C. Fauquet, and D. Lamy, Transmission par la graine du virus du 

clump de l'arachide. Oléagineux, 1978. 33: p. 503-504.

18. ICRISAT, Coordinated research on groundnut rosette virus disease: summary 

proceedings of the consultative group meeting, 8-10 March 1987, Lilongwe, Malawi.

1988.

19. Reddy, D.V.R., Delfosse, P., Lenne, J. M. and Subrahmanyam, P, eds. Groundnut virus 

diseases in Africa: summary and recommendations of the Sixth Meeting of the 

International Working Group, 18-19 Mar 1996, Agricultural Research Council, 

Plant Protection Research Institute, Pretoria, South Africa. 1997. 64.

20. Reddy, D.V.R., B. L. Nolt, H. A. Hobbs, A. S. Reddy, R. Rajeshwar, A. S. Rao, D.D.R. 

Reddy and D. McDonald, Clump virus in India: Isolates, host range, transmission 

and management, in Viruses With Fungal Vectors, J.L. Cooper and M.J.C. Asher, 

Editors. 1988, Assoc. Appl. Biol.: Wellesbourne, UK. p. 239-246.

21. Reddy, D.V.R., M.A. Mayo, and P. Delfosse, Pecluviruses, in Encyclopedia of Virology, 

2nd Ed., R. Webster and A. Gramoff, Editors. 1999, Academic Press: New York.

22. ICRISAT. (International Crops Research Institute for the Semi-Arid Tropics). Viral 

Diseases. [cited 2012 February 17]; Available from: http://www.icrisat.org/btpathology-viral.htm.

23. Baudin, P. and M. Chatenet, Détection sérologieu du PCV isolat canne à sucre, agent de 

la marbrure rouge des feuilles. L'Agronomie Tropicale, 1988. 43: p. 228-234.

24. Dollet, M., C. Fauquet, and J.-C. Thouvenel, Sorghum arundinaceum, a natural host of 
Peanut Clump Virus in Upper Volta. Plant Dis. Rep., 1976. 60: p. 1076-1080.

25. Delfosse, P., A. S. Reddy, A. Legreve, P. S. Devi, K. Thirumala Devi and H. Maraite

D. V. R. Reddy, Indian peanut clump virus (IPCV) infection on wheat and barley: 

symptoms, yield loss and transmission through seed. Plant Pathology, 1999. 48: p. 

273-282.

26. Delfosse, P., A. S. Reddy, K. Thirumala Devi, A. Legrève, J. Risopoulos, D. Doucet, P. 

Shoba Devi, H. Maraite, D. V. R. Reddy, Dynamics of Polymyxa graminis and 

Indian peanut clump virus (IPCV) infection on various monocotyledonous crops and 

groundnut during the rainy season. Plant Pathology, 2002. 51: p. 546-560.
United States Department of Agriculture

Animal and Plant Health Inspection Service

Plant Protection and Quarantine

Plants for Planting Quarantine Pest Evaluation Data Sheet 

January 9th, 2013

In order to prevent the introduction of quarantine pests into the United States, § 319.37-2a allows 

the APHIS Administrator to designate the importation of certain taxa of plants for planting as not 

authorized pending pest risk analysis (NAPPRA). APHIS has determined that the following 

plant taxa should be added to the NAPPRA category. In accordance with paragraph (b)(1) of that 

section, this data sheet details the scientific evidence APHIS evaluated in making the 

determination that the taxa are hosts of a quarantine pest.

Quarantine Pest: Phytophthora kernoviae Brasier Beales & S.A Kirk, sp. Nov.

Hosts: Annona spp., Aesculus spp., Castanea spp., Camellia spp., Drimys spp., Fagus 

spp., Gevuina spp., Hedera spp., Ilex spp., Leucothoe spp., Liriodendron spp., 

Lomatia spp., Magnolia spp. (=Michelia spp.), Pieris spp., Pinus spp., 

Podocarpus spp., Prunus spp., Quercus spp., Rhododendron spp.,

Sequoiadendron spp. (=Sequoia spp.), Vaccinium spp.

__________________________________________________________________________

Taxonomy and description of the pest:

Phytophthora kernoviae is an oomycete pathogen in the Pythiaceae family. Synonyms of 

P. kernoviae include Phytophthora kernovii2, 13 and Phytophthora taxon C12. P. kernoviae 

causes “kernoviae bleeding canker,” “kernoviae dieback,” and “kernoviae leaf blight5.”

Known distribution:

United Kingdom1,4, Ireland7, New Zealand11.

This pest is not known to occur in the United States.

Biology of the pest:

Long distance spread of P. kernoviae is thought to occur primarily by movement of 

infected plant material (e.g., trade)9. Spread may also result from the movement of 
growing media or soil carried on vehicles, machinery, footwear or animals8

. 

P. kernoviae zoospores are contained within sporangia, where they may be aerially or 

splash dispersed4. Mechanisms of zoospore transport from one plant to another include 

rain-splash, wind-driven rain, mist, irrigation or ground water. The infective, swimming 

zoospores penetrate susceptible host material via wounds or natural openings such as 

stomata and lenticels8. After initial penetration, P. kernoviae grows through the plant 

tissue and kills plant cells, which results in visible necrosis. Under suitable conditions, 

asexual reproduction may produce new sporangia, completing P. kernoviae’s life cycle. In 

naturally infected plants, P. kernoviae also produces sexual spores (oospores), which may 

serve as survival structures for the pathogen8. 

On shrub hosts, P. kernoviae causes dieback and leaf blight diseases, with symptoms 

varying slightly among plant species. On Rhododendron spp., early leaf symptoms may 

include a blackening of the leaf tip, the leaf petiole or, in extreme cases, the whole leaf

could fall within a few weeks of infection. Frequently, shoot dieback and cankers occur, 

girdling the stem tissue and causing wilting of the leaves above the lesion8. Infections 

may be found on leaves of any age and on leaves and stems at any height or position on 

the plant. Severe infections result in the death of the whole bush. 

Similar leaf blight symptoms are seen on other shrub hosts. On Michelia doltsopa, 

necrotic leaf tissue is characteristically a dark black-brown color, and infection is 

characterized by drip tip lesions on the leaves that progress along the leaf margins and 

into the tissue of the leaf blade8. Lesions on leaves of Pieris spp. are typically a light tan 

to rusty brown color, with necrosis progressing directly towards and along the midrib 

vein8. Dark necrotic lesions are visible on P. kernoviae-infected leaves of Ilex aquifolium 

(variegated holly) and both leaf infection and stem dieback appear on Prunus 

laurocerasus (cherry laurel)8. On Vaccinium myrtillus (bilberry), early symptoms include 

necrotic black-brown lesions on the leaves and green stems of plants8. Only stem 

infection has been observed on Hedera helix (ivy), to date8.

On tree hosts, P. kernoviae may cause bleeding canker, dieback and/or leaf blight 

diseases. Initial symptoms on Fagus sylvatica (beech) are bleeding lesions on the trunk, 

which may be present from ground level up to 12 m above ground. The bleeding is 

usually dark brown to blue-black, with orange-pink to pink-brown active lesions in the 

inner bark8. Older lesions on beech may appear sunken. Girdling of the entire trunk may 

occur, killing the tree. On Quercus robur (English oak), bleeding lesions are more 

difficult to see because of oak’s thick outer bark ridges and outer bark plates8. Bleeding 

occurs from cracks between the bark ridges, but the thickness of the bark prevents the 

sunken appearance of older cankers. 

Disease symptoms may occur on the foliage, shoots and trunk of Liriodendron tulipifera 

(tulip tree). Internal lesions range from pale brown to blue-black in color, and multiple 
bleeding lesions may be present on the trunk from ground level up to 9 m above ground8

. 

Leaf lesions are mostly restricted to tips and margins and appear dark black in color. 

Shoot dieback also occurs, and infected shoots are defoliated8. No evidence of sunken or 

bleeding cankers has been observed on Quercus ilex (holly oak), but P. kernoviae does 

cause severe leaf necrosis and dieback of Q. ilex epicormic shoots8.

On Magnolia spp., P. kernoviae infection occurs anywhere on the leaf surface. Multiple 

leaf infections may appear as numerous dark brown necrotic patches, giving leaves a 

spotty appearance8. The necrotic spots may merge and develop towards the leaf midrib. 

Well-developed lesions appear as mottling, which may have angular edges8. Uninfected 

tissue between necrotic areas becomes chlorotic. Petioles may become infected leading to 

disease progression along the leaf base. Infected buds appear light khaki grey in color8. 

Damage potential of pest:

In England and Wales, between October 2003 and January 2008, there were 52 outbreaks 

of P. kernoviae infection in non-nursery locations5. All but one of these outbreaks are 

subject to ongoing eradication or containment action. In 2004, the damage potential of P. 

kernoviae was considered sufficient to warrant a statutory order in England that prohibits 

the removal of host plants or parts of host plants from a designated area without 

inspection and written authorization2. P. kernoviae has the potential to attack a wide 

range of important forest trees and could cause significant damage to U.S. agriculture and 

the environment. 

Control:

At present, no specific control measures for P. kernoviae have been identified. The 

current management strategy is the utilization of good cultural practices

8

. Susceptible 

plants should be monitored for symptoms. Shears and tools should be regularly cleaned 

with an appropriate product. Plants with wounded leaves should be handled carefully, as a 

wounded leaf may be more susceptible to infection8. Susceptible host plants should be 

pruned in dry weather. If possible, plants should be watered in the morning, and potted 

plants should not be subjected to standing in water for any length of time. Plants should 

be spaced appropriately to allow for good air movement. 

Known host range:

Annona spp.

3, 5

, Aesculus spp.5, Castanea spp.5, Camellia spp.

4

, Drimys spp.5, 9, Fagus 

spp.4, 5, Gevuina spp.4, 5, Hedera spp.5, 9, Ilex spp.5, 9, Leucothoe spp.5, 

Liriodendron spp.4, 5, Lomatia spp.9, Magnolia spp.5, 9 (=Michelia spp.4, 5) , Pieris spp.

4,5,9

, Pinus spp.5, Podocarpus spp.5, 9, Prunus spp.5,9, Quercus spp.

4,5,6

, Rhododendron 

spp.4, 8,10, Sequoiadendron spp.9(=Sequoia spp.), Vaccinium spp.5, 9.
Action under NAPPRA:

The importation of the following genera of plants for planting, excluding seeds, that are 

hosts of Phytophthora kernoviae, is not authorized pending a pest risk analysis 

(NAPPRA) from all countries:

Camellia, Drimys, Gevuina, Lomatia, Sequoiadendron (Sequoia).

The importation of the following plants for planting genera, excluding seeds that are 

hosts of Phytophthora kernoviae, are not authorized pending a pest risk analysis 

(NAPPRA) from all countries, except those listed after the plant genus:

Annona Costa Rica 

Fagus Canada

Hedera Canada, Colombia, Costa Rica, Guatemala, 

Israel, and Mexico

Ilex Canada

Leucothoe Canada

Liriodendron Canada

Magnolia (Michelia) Canada, China, and South Africa

Pieris Canada

Podocarpus Canada, China

Quercus Canada

Rhododendron Canada, Japan

Vaccinium Australia, Canada.

The importation of the following plants for planting genera that are hosts of Phytophthora 

kernoviae is currently regulated under 7 CFR 319.37 and these genera are therefore not 

included, at this time, as potential candidates for listing on this NAPPRA data sheet:

Aesculus, Castanea. 

References:

1. Anonymous. Plant disease found in Scotland. The Scottish Government News. Accessed 

10 Jan 2008 at http://www.scotland.gov.uk/News/Releases/2008/01/10144052

2. Anonymous. The plant health (Phytophthora kernovii Management Zone)(England) order 

2004. Statutory Instrument 2004 No. 3367.

3. Braithwaite, M., M. S. Bullians, J. M. Pay, G. S. C. Gill, and C. F. Hill. 2007. A disease 

survey of cherimoya orchards in Northland, New Zealand. Proceedings of the 16th 
Biennial Australasian Plant Pathology Conference, Adelaide: Australasian Plant 

Pathology Society

4. Brasier, C. M., P. A. Beales, S. A. Kirk, S. Denman, and J. Rose. 2005. Phytophthora 

kernoviae sp. nov., an invasive pathogen causing bleeding stem lesions on forest 

trees and foliar necrosis of ornamentals in the UK. Mycol Res. 109:853-859

5. CSL. Revised Summary Pest Risk Analysis for Phytophthora kernoviae. Accessed 24 Feb 

2012 at 

http://www.fera.defra.gov.uk/plants/plantHealth/pestsDiseases/phytophthora/pKerno

viae/

6. Denman, S., S. A. Kirk, E. Moralejo, and J. F. Webber. 2009. Phytophthora ramorum

and Phytophthora kernoviae on naturally infected asymptomatic foliage. EPPO 

Bulletin. 39:105-111

7. EPPO. First report of Phytophthora kernoviae in Ireland (2010/148). EPPO Reporting 

Service. Vol. 9. Accessed 24 Feb 2012 at 

http://archives.eppo.org/EPPOReporting/2010/Rse-1009.pdf

8. FERA. A new threat to our trees and woodlands: Phytophthora kernoviae disease 

factsheet. Accessed 25 Feb 2012 at www.fera.defra.gov.uk/pKernoviaeFactsheet

9. FERA. Phytophthora kernoviae - FAQs. Accessed 24 Feb 2012 at 

http://www.fera.defra.gov.uk/plants/plantHealth/pestsDiseases/phytophthora/pKerno

viae/faqs.cfm

10. Fichtner, E. J., D. M. Rizzo, S. A. Kirk, and J. F. Webber. 2011. Infectivity and 

sporulation potential of Phytophthora kernoviae to select North American native 

plants. Plant Pathol.10.1111/j.1365-3059.2011.02506.x

11. Goheen, E. M., and S. J. Frankel (ed.). 2009. Proceedings of the fourth meeting of the 

International Union of Forest Research Organizations (IUFRO) Working Party 

S07.02.09: Phytophthoras in forest and natural ecosystems. Gen. Tech. Rep. PSWGTR-221. Albany, CA: U.S. Department of Agriculture, Forest Service, Pacific 

Southwest Research Station. pp. 47-53.

12. Hughes, K. J. D., R. Griffin, N. Boonham, and A. J. Inman. 2005. Development of 

molecular diagnostics for Phytophthora taxon C a new Phytophthora threatening UK 

trees, woodlands and ornamental plants. Sudden Oak Death Science Symposium II, 

18-21 January 2005, Monterey, CA

13. Morton, C. Native oaks susceptible to new tree disease. Forestry Commission News. 9 

Nov 2004
United States Department of Agriculture

Animal and Plant Health Inspection Service

Plant Protection and Quarantine

Plants for Planting Quarantine Pest Evaluation Data Sheet 

January 9th, 2013

In order to prevent the introduction of quarantine pests into the United States, § 319.37-2a allows 

the APHIS Administrator to designate the importation of certain taxa of plants for planting as not 

authorized pending pest risk analysis (NAPPRA). APHIS has determined that the following 

plant taxa should be added to the NAPPRA category. In accordance with paragraph (b)(1) of that 

section, this data sheet details the scientific evidence APHIS evaluated in making the 

determination that the taxa are hosts of a quarantine pest.

Quarantine Pest: Puccinia buxi Sowerby 1809

Hosts: Buxus spp.

4, 5

__________________________________________________________________________

Taxonomy and description of the pest:

Puccinia buxi Sowerby 1809

Urediniomyces: Uredinales (Order: Family)

Common name of disease – boxwood rust

Known distribution:

Japan, China, France, Georgia, Germany, Greece, Ireland, Italy, Poland, Portugal, 

Spain, Switzerland, and the United Kingdom4. On May 2006 boxwood plants at a 

facility in Pennsylvania where found infected with Puccinia buxi. These plants 

were earlier imported from Greece8. This infestation was subsequently eradicated. 

Biology of the pest:

Puccinia buxi is a rust fungus with two spore forms which completes its life cycle on one 

host, Buxus sp.6, 9. The teliospores, a natural resting stage, develop within host tissues 

during the winter and break through the plant’s epidermis in the spring10. Teliospores 
germinate in spring and early summer to produce basidia, then infective basidiospores 

which can infect new hosts or new leaves on the same host plant. Infections develop 

slowly, with lesions expanding in the late summer and fall, which appear initially as 

thickenings of the leaf10. Buxus spp. plants growing in shaded and moist conditions 

appear to be more susceptible (or conditions are more suitable for infection), than plants 

grown in full sun or on south-facing slopes, and leaf age also appears to impact 

susceptibility3. 

Damage potential of pest:

Although not widespread by natural means, it does appear to be readily transported on 

nursery stock. It may be difficult to identify and detect during port of entry inspection and 

therefore likely to be unintentionally transported internationally. Infected Buxus spp. may 

show blister-like pustules forming on infected leaves as the telia develop8, although there 

may also be a latent stage prior to symptom expression. Since 2003, APHIS Plant 

Inspection Stations have recorded 10 interceptions of P. buxi on Buxus spp. for decorative 

purpose and 3 interceptions on Buxus spp. for propagation from various countries. 

Control: 

Cultural control measures, such as growing plant material in a greenhouse using sanitary 

procedures, including inspections, can be useful to control Puccinia spp. in ornamental 

production. For example, heat treatments can kill rust spores and eradicate the pathogen

in plant material; geranium rust (Puccinia pelargoni-zonalis) can be eradicated by hot 

water treatment of cuttings, although damage to the host can occur7. For most rusts, water 

is necessary for infection, so overhead irrigation should be avoided. Infected leaves with 

teliospores should be removed, rapid composted, or buried to stop the life cycle, and 

prevent further infections. There are several fungicides that can be used to protect plants 

from infection7. 

Known host range:

Buxus spp.4, 5

Action under NAPPRA:

The importation of Buxus spp. plants for planting, excluding seed but including cut 

flowers and greenery, hosts of Puccinia buxi, is not authorized pending pest risk analysis 

(NAPPRA) from all countries except Canada. 
References:

1. 7CFR § 319.37. 2011. Code of Federal Regulations, Title 7, Part 319.37, (7 CFR §319 -

Foreign quarantine notices: subpart—Plants for Planting). United States 

Government.

2. Agriculture Quarantine Activity System Databases. 2011. Pest Identification Database 

(PestID).

3. Durrieu, G. (2001). More about box rust (Puccinia buxi). Mycologist, 15 (3), 144-145.

4. Farr, D.F., and Rossman, A.Y. 2011. Fungal Databases, Systematic Mycology and 

Microbiology Laboratory, ARS, USDA. Accessed March 5, 2012, online at 

http://nt.ars-grin.gov/fungaldatabases/index.cfm

5. Germplasm Resources Information Network - (GRIN) [Online Database]. USDA, ARS, 

National Genetic Resources Program. National Germplasm Resources Laboratory, 

Beltsville, Maryland. Last accessed March 5, 2012, online at http://www.arsgrin.gov/cgi-bin/npgs/html/splist.pl?1826

6. Grove, W. B. (1913). The British rust fungi (Uredinales): their biology and classification. 

Cambridge University Press, Cambridge. 

7. Koike, S. T., Wilen, C. A., Raabe, R. D., McCain, A. H. and Grebus, M. E. 2009. UC 

IPM Pest Management Guidelines: Floriculture and Ornamental Nurseries UC ANR 

Publication 3392

8. Phytosanitary Alert System (PAS), NAPPO. 2006. Boxwood rust (Puccinia buxi) 

incident in Pennsylvania. Accessed March 5, 2012 online at 

http://www.pestalert.org/oprDetail.cfm?oprID=202

9. Preece, T. F. 2000. The strange story of box rust, Puccinia buxi, in Britain. Mycologist, 

14 (3), 104-106. 

10. Wilson, M., and D. M. Henderson. (1966). British rust fungi. Cambridge University 

Press, Cambridge. 

11. Yun, H.Y. Systematic Mycology and Microbiology Laboratory, ARS, USDA. Invasive 

Fungi. Box Rust or Boxwood Rust - Puccinia buxi. Retrieved March 5, 2012 , online 

at http://nt.ars-grin.gov/taxadescriptions/factsheets/index.cfm?thisapp=Pucciniabuxi