Petition for the Determination of Nonregulated Status for 

Herbicide-Tolerant 73496 Canola 

We submit this petition under 7 CFR §340.6 to request that the Administrator, Animal and Plant Health 

Inspection Service, make a determination that the article should no longer be regulated under 

7 CFR §340. 

Submitting Company: 

Pioneer Hi-Bred International, Inc. 

7100 NW 62nd Avenue 

PO Box 1014 

Johnston, IA 50131-1014 

Submitted by: 

Natalie Weber, Registration Manager 

Pioneer Hi-Bred International, Inc. 

DuPont Agricultural Biotechnology 

DuPont Experimental Station, E353/305C 

PO Box 80353 

Wilmington DE 19880-0353 

Telephone: 302-695-8160 

Fax: 302-695-3075 

Prepared by: 

John Larsen, Natalie Weber, Jennifer Tallman, Donald MacKenzie, Alejandra Pascual, Keri Carstens, 

Kate Hayter, Kara Califf, Kent Brink, Theresa Sult, and Holly Hettinger

OECD Unique Identifier: DP-Ø73496-4 

Submission date: February 25, 2011 

Revised submission date: August 3, 2011 
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73496 Canola 

TABLE OF CONTENTS 

 

LIST OF TABLES ............................................................................................................................................... 5 

LIST OF FIGURES ............................................................................................................................................. 7 

RELEASE OF INFORMATION .............................................................................................................................. 9 

CERTIFICATION .............................................................................................................................................. 10 

SUMMARY ..................................................................................................................................................... 11 

ABBREVIATIONS, ACRONYMS, AND DEFINITIONS .............................................................................................. 13

I. RATIONALE FOR THE DEVELOPMENT OF 73496 CANOLA ............................................................................. 15 

I-A. Basis for the Request for a Determination of Nonregulated Status under 7 CFR §340.6 .................. 15 

I-B. Benefits of 73496 Canola .................................................................................................................... 15 

I-C. Submissions to Other Regulatory Agencies ........................................................................................ 15 

II. THE BIOLOGY OF CANOLA ......................................................................................................................... 16 

II-A. Canola as a Crop ................................................................................................................................ 16 

II-B. Description of the Non-Transformed Recipient Canola Line .............................................................. 16 

III. METHOD OF DEVELOPMENT OF 73496 CANOLA ........................................................................................ 16 

III-A. Description of Transformation, Selection, and Breeding .................................................................... 16 

III-B. Selection of Comparators for 73496 Canola ..................................................................................... 18

IV. DONOR GENES AND REGULATORY SEQUENCES ....................................................................................... 18 

IV-A. Fragment PHP28181A Used in Transformation ............................................................................... 18 

IV-B. Identity and Source of the gat4621 Gene Cassette on Fragment PHP28181A ............................... 20 

V. GENETIC CHARACTERIZATION OF 73496 CANOLA ..................................................................................... 21 

V-A. Molecular Analysis Overview ............................................................................................................. 21 

V-B. Results of Molecular Characterization of 73496 Canola by Southern Blot Analysis ......................... 21 

V-C. Inheritance and Genetic Stability of the Introduced Trait in 73496 Canola ....................................... 34 

V-D. Conclusions on Molecular Characterization and Genetic Stability of 73496 Canola ........................ 35 

VI. CHARACTERIZATION OF THE INTRODUCED GAT4621 PROTEIN ................................................................. 36 

VI-A. The GAT4621 Protein ....................................................................................................................... 36

VI-B. Physicochemical Characterization of the GAT4621 Protein ............................................................. 38

VI-C. Concentrations of GAT4621 in 73496 Canola .................................................................................. 38 

VI-D. Summary of the Food and Feed Safety Assessment for the GAT4621 Protein ............................... 40 

VII. AGRONOMIC PERFORMANCE AND ECOLOGICAL OBSERVATIONS .............................................................. 42 

VII-A. Germination Evaluations .................................................................................................................. 42 

VII-B. Field Trial Evaluations ...................................................................................................................... 43 

VII-C. Ecological Observations .................................................................................................................. 52 

VII-D. Conclusions on Agronomic Performance and Ecological Observations ......................................... 53
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TABLE OF CONTENTS (CONTINUED) 

VIII. COMPOSITIONAL ASSESSMENT .............................................................................................................. 54

VIII-A. Proximate and Fiber Analysis ......................................................................................................... 55 

VIII-B. Fatty Acid Analysis .......................................................................................................................... 57

VIII-C. Amino Acid Analysis ....................................................................................................................... 60

VIII-D. Vitamin Analysis ............................................................................................................................... 63

VIII-E. Mineral Analysis ............................................................................................................................... 65

VIII-F. Glucosinolate Analysis .................................................................................................................... 67

VIII-G. Secondary Metabolite and Anti-Nutrient Analysis ........................................................................... 70

VIII-H. Acetylated Amino Acids .................................................................................................................. 72

VIII-I. Conclusions on the Compositional Analysis of 73496 Canola ......................................................... 75

IX. ASSESSMENT OF PLANT PEST RISK FOR 73496 CANOLA ......................................................................... 77

IX-A. Environmental Assessment of the Transgenic Protein ..................................................................... 77 

IX-B. Fate of Transgenic DNA in Humans and Animals ............................................................................ 77 

IX-C. Weediness Potential of 73496 Canola .............................................................................................. 77 

IX-D. Gene Flow Assessment .................................................................................................................... 81 

APPENDIX 1. MATERIALS AND METHODS FOR GENETIC CHARACTERIZATION OF 73496 CANOLA ..................... 92 

1.1. Southern Blot Characterization of 73496 Canola ............................................................................... 92 

1.2. Mendelian Inheritance and Genetic Stability of the Trait in 73496 Canola ......................................... 95 

APPENDIX 2. DESCRIPTION OF STATISTICAL ANALYSES PERFORMED FOR AGRONOMIC AND NUTRIENT 

COMPOSITION DATA ...................................................................................................................................... 97

2.1. Statistical Analysis of Germination Data ............................................................................................. 97 

2.2. Statistical Analyses of Agronomic Characteristic, Yield, and Nutrient Composition Data .................. 97 

2.3. Special Considerations ..................................................................................................................... 100 

2.4. SAS Procedures ................................................................................................................................ 101 

APPENDIX 3. MATERIALS AND METHODS FOR DETERMINATION OF GAT4621 PROTEIN CONCENTRATIONS..... 102 

3.1. Plant Material .................................................................................................................................... 102 

3.2. Storage and Processing of ELISA Samples ..................................................................................... 102 

3.3. Protein Extraction from Ground Canola Tissues .............................................................................. 102 

3.4. Determination of GAT4621 Protein Concentrations ......................................................................... 102 

APPENDIX 4. CHARACTERIZATION OF THE PLANT-EXPRESSED AND MICROBIAL-EXPRESSED 

GAT4621 PROTEINS ................................................................................................................................... 104 

APPENDIX 5. ECOLOGICAL OBSERVATIONS FOR 73496 CANOLA .................................................................. 111 

APPENDIX 6. USDA AND CANADIAN FIELD TRIALS OF 73496 CANOLA ......................................................... 115 

APPENDIX 7. SURVEY OF POTENTIAL SUBSTRATES FOR GAT4621 .............................................................. 116 
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TABLE OF CONTENTS (CONTINUED) 

APPENDIX 8. SAFETY AND HISTORY OF CONSUMPTION OF ACETYLATED AMINO ACIDS.................................. 118 

8.1. Summary ........................................................................................................................................... 118 

8.2. Function and Use of Acetylated Amino Acids ................................................................................... 118 

8.3. History of Safe Use of Acetylated Amino Acids ................................................................................ 118 

8.4. Presence of NAA, NAG, NAT, NAS, and NAGly in Biological Systems ........................................... 119 

8.5. History of Safe Consumption of NAA, NAG, NAS, NAT, and NAGly ................................................ 121 

8.6 Toxicology Studies Conducted with NAA, NAG, NAS, NAT, and NAGly .......................................... 121 

APPENDIX 9. HERBICIDE RESISTANT WEEDS ............................................................................................... 122 

9.1. Evolution of Herbicide Resistant Weeds ........................................................................................... 122 

9.2. Characteristics of Glyphosate ........................................................................................................... 124 

9.3. Evolution of Resistance to Glyphosate ............................................................................................. 124 

9.4. Stewardship of Herbicide Tolerant 73496 Canola ............................................................................ 125 

APPENDIX 10. GAT4621 NEW PROTEIN CONSULTATION ............................................................................. 130 

REFERENCES .............................................................................................................................................. 148
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73496 Canola 

LIST OF TABLES 

Table 1. Generations and Comparators Used for Analysis of 73496 Canola ........................................... 18 

Table 2. Description of the Genetic Elements in Fragment PHP28181A .................................................. 20 

Table 3. Description of DNA Probes Used for Southern Blot Hybridization .............................................. 24 

Table 4. Summary of Expected and Observed Hybridization Fragments on Southern 

Blots for 73496 Canola ............................................................................................................... 25 

Table 5. Summary of Genotypic and Phenotypic Results for 73496 Canola ............................................ 35 

Table 6. Comparison of Amino Acid Sequence Similarity Between Parental GAT 

Enzymes and GAT4621 .............................................................................................................. 38 

Table 7. Concentrations of GAT4621 Protein in 73496 Canola Plant Tissues ......................................... 39 

Table 8. Description of Seed Germination Conditions .............................................................................. 42 

Table 9. Summary of Germination Results for 73496 Canola ................................................................... 43 

Table 10. Site Locations for 2008 and 2009 Field Trial Experiments ........................................................ 45 

Table 11. Experiment A: 2008 Field Trial Agronomic Characteristics Measured ..................................... 47 

Table 12. Experiment A: Summary of Agronomic Characteristics of 73496 Canola 

Across Seven Locations ............................................................................................................ 48 

Table 13. Experiment B: 2009 Field Trial Agronomic Characteristics Measured ..................................... 49 

Table 14. Experiment B: Summary of Agronomic Characteristics of 73496 Canola 

Across Three Locations ............................................................................................................ 51

Table 15. Summary Analysis of Proximate and Fiber Results in Canola Seed ........................................ 56 

Table 16. Summary Analysis of Fatty Acid Composition in Canola Seed ................................................. 58 

Table 17. Summary Analysis of Amino Acid Composition in Canola Seed ............................................... 61 

Table 18. Summary Analysis of Vitamin Composition in Canola Seed ..................................................... 64 

Table 19. Summary Analysis of Mineral Composition in Canola Seed ..................................................... 66 

Table 20. Summary Analysis of Glucosinolate Composition in Canola Seed ........................................... 68 

Table 21. Summary Analysis of Secondary Metabolite and Anti-Nutrient Composition 

in Canola Seed .......................................................................................................................... 71

Table 22. Concentrations of Acetylated Amino Acids in Canola Seed ...................................................... 73 

Table 23. Concentrations of Acetylated Amino Acids in Canola Whole Plant Samples ........................... 74 
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73496 Canola 

LIST OF TABLES (CONTINUED) 

Table 24. Concentrations of Acetylated Amino Acids in Processed Fractions Produced 

from 73496 and Control Canola ................................................................................................ 75 

Table 25. Common Characteristics of Weeds ........................................................................................... 78 

Table 26. List of Species with Reports of Hand-Pollinated or Spontaneous Hybridization 

with B. napus ............................................................................................................................. 84

Table 5.1. United States: Insect Stressor Comparison between 73496 Canola and Controls ............... 111 

Table 5.2. United States: Disease Stressor Comparison between 73496 Canola and Controls ........... 112 

Table 5.3. Canada: Insect Stressor Comparison between 73496 Canola and Controls ........................ 113 

Table 5.4. Canada: Disease Stressor Comparison between 73496 Canola and Controls .................... 114 

Table 6.1. USDA Field Trials of 73496 Canola........................................................................................ 115 

Table 6.2. Canada Field Trials of 73496 Canola ..................................................................................... 115 

Table 7.1. kcat/KM for the GAT4621 Enzyme on Glyphosate and Selected Amino Acids ........................ 116 

Table 8.1. Full list of Studies Conducted with NAA, NAG, NAS, NAT, and NAGly ................................. 121 

Table 9.1. Herbicide Resistant Weeds Summary Table (Heap, 2010b) ................................................. 123 
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73496 Canola 

LIST OF FIGURES

Figure 1. Schematic Diagram of the Development of 73496 Canola ........................................................ 17 

Figure 2. Breeding Diagram for 73496 Canola and Generations Used for Analyses ............................... 17 

Figure 3. Schematic Diagram of Fragment PHP28181A .......................................................................... 19 

Figure 4. Schematic Diagram of Plasmid PHP28181 ............................................................................... 19 

Figure 5. Restriction Enzyme Map of PHP28181A DNA Fragment .......................................................... 25 

Figure 6. Restriction Enzyme Map of Plasmid PHP28181 ........................................................................ 26 

Figure 7. Physical Map of the Insertion in 73496 Canola ......................................................................... 26 

Figure 8. Southern Blot Analysis of 73496 Canola: Nco I Digestion and UBQ10 

Promoter Probe ......................................................................................................................... 27 

 

Figure 9. Southern Blot Analysis of 73496 Canola: Nco I Digestion and gat4621 Probe ......................... 28 

Figure 10. Southern Blot Analysis of 73496 Canola: Nco I Digestion and pinII Terminator 

Probe ....................................................................................................................................... 29 

Figure 11. Southern Blot Analysis of 73496 Canola: Ssp I Digestion and UBQ10 

Promoter Probe ....................................................................................................................... 30 

Figure 12. Southern Blot Analysis of 73496 Canola: Ssp I Digestion and gat4621 Probe ....................... 31 

Figure 13. Southern Blot Analysis of 73496 Canola: Ssp I Digestion and pinII 

Terminator Probe ..................................................................................................................... 32 

Figure 14. Southern Blot Analysis of 73496 Canola: Nco I Digestion and Plasmid 

Backbone Probe ..................................................................................................................... 33 

Figure 15. Deduced Amino Acid Sequence of the GAT4621 Protein ....................................................... 36 

Figure 16. Enzymatic Activity of the GAT4621 Protein ............................................................................. 37 

Figure 17. Map of Locations for Agronomic Data Collection for 73496 Canola ........................................ 45 

Figure 18. Map Indicating Zones with Similar Growing Conditions for Field Trials ................................... 46 

Figure 19. Production Climate Zones for Spring, Biennial, and Winter Canola ........................................ 82 

Figure 20. Genome Relationships Among Cultivated Brassicaceae ........................................................ 83 

Figure 4.1. SDS-PAGE Analysis of the Plant-Expressed GAT4621 Protein .......................................... 104 

Figure 4.2. Western Immunoblot Analysis of the Plant-Expressed and 

Microbial-Expressed GAT4621 Proteins .............................................................................. 105 

Figure 4.3. N-Terminal Amino Acid Sequence Analysis of the Plant-Expressed 

and Microbial-Expressed GAT4621 Proteins ....................................................................... 106 
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73496 Canola 

LIST OF FIGURES (CONTINUED) 

Figure 4.4. Deduced GAT4621 Amino Acid Sequence Showing Matching Peptides 

from MALDI-MS ................................................................................................................... 106 

Figure 4.5. Glycosylation Analysis of the Plant-Expressed GAT4621 Protein ........................................ 107 

Figure 9.1. Number of Resistant Biotypes of Weeds by Herbicide Class and Year (Heap, 2010d) ....... 124 

Figure 9.2. Resistant Weed Management Label ..................................................................................... 127
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73496 Canola 

Release of Information 

Pioneer Hi-Bred International, Inc. (Pioneer) is submitting the information in this assessment for review by 

USDA as part of the regulatory process. By submitting this information, Pioneer does not authorize its 

release to any third party except to the extent it is requested under the Freedom of Information Act 

(FOIA), 5 U.S.C., Section 552; USDA complies with the provisions of FOIA and USDA’s implementation 

regulations (7 CFR Part 1.4); and this informaiton is responsive to the specific request. Except in 

accordance with the Freedom of Information Act, Pioneer does not authorize the release, publication or 

other distribution of this information (including website posting) without Pioneer’s prior notice or consent. 

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73496 Canola 

Summary 

Pioneer Hi-Bred International, Inc. (Pioneer) is submitting this Petition for Determination of Nonregulated 

Status for herbicide-tolerant canola event DP-Ø73496-4, hereafter referred to as 73496 canola. Canola 

line 73496 was developed by Pioneer Hi-Bred, a DuPont Business. The data in this petition support the 

conclusion that 73496 canola is not likely to pose an increased plant pest potential. Therefore, Pioneer 

requests a determination from USDA - Animal and Plant Health Inspection Service (APHIS) that 73496 

canola and any crosses of this line with other nonregulated Brassica napus no longer be considered 

regulated articles under 7 CFR §340. 

73496 canola is a transgenic plant line that is tolerant to the herbicidal active ingredient glyphosate. The 

availability of 73496 canola will provide an alternative to currently available glyphosate-tolerant canola 

lines. Herbicide-tolerant 73496 canola will provide the same benefits as currently available glyphosatetolerant canola lines, in that growers will be able to proactively manage weed populations and delay the 

development of adverse populations of troublesome weeds. 

73496 canola has been genetically modified to express the GAT4621 (glyphosate acetyltransferase) 

protein. The gat4621 gene is a variant of three gat genes from the common soil bacterium Bacillus 

licheniformis. The GAT4621 protein is equivalent to the protein expressed in 98140 maize deregulated 

by USDA in 2009 and reviewed by FDA in 2008, and is encoded by the gat4621 gene which confers 

tolerance to glyphosate-containing herbicides by acetylating glyphosate and thereby rendering it nonphytotoxic. In 73496 canola, the expression of the gat4621 gene is driven by the Arabidopsis

polyubiquitin constitutive promoter. 

73496 canola was generated using biolistic transformation of canola microspores with a gel-purified DNA 

fragment isolated from plasmid PHP28181 containing the gat4621 gene cassette. Molecular 

characterization of 73496 canola by Southern blot analysis confirmed that a single, intact PHP28181A 

DNA fragment has been inserted into the genome with no plasmid backbone DNA. Segregation analysis 

of 73496 canola confirmed Mendelian inheritance of the gat4621 gene. 

The potential for allergenicity and toxicity of 73496 canola was evaluated by examining the allergenic 

potential of canola as a crop and by assessing the allergenic and toxic potential of the GAT4621 protein. 

Canola is not a common allergenic food and the modification in 73496 canola is not expected to alter the 

allergenic potential of canola. The allergenic potential of the GAT4621 protein was assessed using a 

weight-of-evidence approach using guidance from the Codex Alimentarius Commission. Bioinformatic 

analyses revealed no biologically significant identities to known or putative protein allergens or toxins for 

the GAT4621 protein sequences. The GAT4621 protein is not glycosylated and is rapidly digested (within 

30 seconds) in simulated gastric fluid. In simulated intestinal fluid, the GAT4621 protein hydrolyzed within 

5 minutes. There was no evidence of acute toxicity in mice for GAT4621 at a dose of 1640 mg protein 

per kg of body weight. Based on the GAT4621 protein levels in 73496, exposure levels would be 

exponentially lower than the tested doses. These data support the conclusion that the GAT4621 protein 

is unlikely to cause an allergic reaction in humans or be a toxin in humans or animals and therefore 

support the food and feed safety of GAT4621. A New Protein Consultation for the GAT4621 protein was 

submitted to FDA on January 31, 2007 and completed on October 7, 2009. 

73496 canola has been field tested since 2007 in the major canola-growing regions of Canada and the 

United States. All field tests that have occurred in the United States were under field permits and 

notifications granted by USDA - APHIS. Comprehensive agronomic performance for 73496 canola were 

conducted in replicated field studies at a total of 10 locations across Canada and the United States. 

Characteristics such as early growth, days to flower, days to maturity, plant height, yield, seedling vigor, 

flowering duration, lodging, shattering, disease incidence and insect damage were assessed. Seed 

germination and dormancy data were also collected in laboratory experiments. All field trials of 73496 

canola were observed for naturally occurring disease or insect biotic stressors. Analysis of agronomic 

and ecological data showed no biologically meaningful differences between 73496 canola and control 

canola lines, indicating the similarity of 73496 canola to conventional canola and indicating no plant pest 
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73496 Canola 

characteristics or selective advantage in natural habitats. These data support the conclusion of 

agronomic comparability of 73496 canola to commercially available canola with respect to the lack of 

increased weediness and plant pest potential. 

Extensive nutrient composition analysis of seed was conducted to compare the composition of 73496 

canola to that of a control line and five commercial canola varieties. Compositional analysis of 73496 

canola was used to evaluate any changes in the levels of key nutrients, anti-nutrients, and secondary 

metabolites in accordance with the OECD consensus document for new varieties of canola. Based on 

the results of the compositional evaluation, the seed of 73496 canola are comparable to commercially 

available canola. Along with the agronomic data included in this petition, compositional comparability is a 

general indicator that 73496 canola will not exhibit unintended effects due to the inserted DNA. 

In addition to glyphosate, the GAT4621 protein is known to acetylate certain free amino acids (Laspartate, L-glutamate, glycine, L-serine, and L-threonine) resulting in the production of N-acetylaspartate 

(NAA), N-acetylglutamate (NAG), N-acetylglycine (NAGly), N-acetylserine (NAS), and N-acetylthreonine 

(NAT). The efficiency of acetylation of free amino acids by GAT proteins is considerably lower than the 

activity displayed toward glyphosate. Because of the potential for these five acetylated amino acids to be 

increased in 73496 canola, concentrations were measured in seed, whole plant, and processed product 

samples. An increase in concentrations of NAA, NAG, and, in some instances, NAGly, NAS, and NAT, 

was confirmed through the analysis of seed, whole plant, and processed product samples. Low but 

quantifiable concentrations of each N-acetylated amino acid were found in each sample type except 

refined, bleached, deodorized (RBD) oil, where levels were either not detectable or below the limit of 

quantification. These five acetylated amino acids are not novel substances as they are present in 

conventional canola as well as in other plants. 

In conclusion, based on the data contained herein that support the conclusion that 73496 canola is not 

likely to pose an increased plant pest potential, Pioneer requests that APHIS grant the request for a 

determination of nonregulated status for 73496 canola and any crosses of this line with other 

nonregulated Brassica napus. 
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73496 Canola 

Abbreviations, Acronyms, and Definitions 

~ approximately 

1822B canola donor line used for transformation 

1822R non-transgenic canola line used in breeding 

5300B non-transgenic canola line used in breeding 

5536F non-transgenic canola line used in breeding 

6393B non-transgenic canola line used in breeding 

6395B non-transgenic canola line used in breeding 

73496 canola canola lines containing the DP-Ø73496-4 event 

ACY1 Aminoacylase I 

ADF acid detergent fiber 

ALS Acetylactate synthase 

AOSA Association of Official Seed Analysis 

APHIS Animal and Plant Health Inspection Service of USDA 

ATCC American Type Culture Collection 

BamH I Restriction enzyme from Bacillus amyloliquefaciens

BAR phosphinothricin acetyltransferase 

BBCH Bundesanstalt, Bundessortenamt and Chemical industry 

bla (ApR) Ampicillin resistance gene 

B. licheniformis Bacillus licheniformis 

B. napus Brassica napus L.

bp base pair 

CAC Codex Alimentarius Commission 

CCA Certified Crop Adviser 

CFIA Canadian Food Inspection Agency 

CFR Code of Federal Regulations 

CNS Central nervous system 

CPSI Carbamyl phosphate synthetase I 

CT threshold cycle 

Da Dalton 

df degrees of freedom 

DIG digoxygenin 

DP-Ø73496-4 OECD identifier for herbicide-tolerant canola event

DP-Ø9814Ø-6 OECD identifier for approved herbicide-tolerant maize event

DNA deoxyribonucleic acid 

E. coli Escherichia coli 

E score expectation score 

ELISA enzyme linked immunosorbent assay 

EPA Environmental Protection Agency 

EPSPS enolpyruvylshikimate-3-phosphate synthase 

EU European Union 

FARRP Food Allergy Research and Resource Program 

FDA Food and Drug Administration 

FDR false discovery rate 

FOIA Freedom of Information Act 

GAT glyphosate acetyltransferase 

GAT4621 specific GAT protein 

gat4621 specific gat gene 

GLMM Generalized Linear Mixed Model 

GNAT GCN 5-related N-acetyltransferases 

HaeIII Restriction enzyme from Haemophilus aegyptius

Hind III Restriction enzyme from Haemophilus influenzae
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73496 Canola 

Abbreviations, Acronyms, and Definitions (continued) 

HRAC Herbicide Resistance Action Committee 

HRP horseradish peroxidase 

IgG immunoglobulin G 

ISAAA International Service for the Acquisition of Agri-biotech Applications 

kb kilobase pair 

kDa kilodalton 

LLOQ lower limit of quantitation 

LS-Mean Least Squares Mean 

MALDI-MS matrix assisted laser desorption ionization mass spectrometry 

mRNA messenger ribonucleic acid 

NAA N-acetylaspartate 

NAG N-acetylglutamate 

NAGly N-acetylglycine 

NAGS N-acetylglutamate synthase

NAS N-acetylserine 

NAT Nacetylthreonine 

NCBI National Center for Biotechnology Information 

Nco I Restriction enzyme from Nocardia corallina

NDF neutral detergent fiber 

Not I Restriction enzyme from Nocardia otitidis-caviarum

NRCS Natural Resources Conservation Service 

OD optical density 

OECD Organisation for Economic Co-operation and Development 

OGTR Office of the Gene Technology Regulator 

PAT phosphinothricin acetyltransferase 

PCR polymerase chain reaction 

PEP phosphoenolpyruvate 

pinII proteinase inhibitor II 

PMRA Pest Management Regulatory Agency (Canada) 

ppm parts per million 

REML Residual Maximum Likelihood 

SAS Statistical Analysis Software 

SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis 

Ssp I Restriction enzyme from Sphaerotilus species 

UBQ10 polyubiquitin gene from Arabidopsis thaliana

USDA United States Department of Agriculture 

USDA-ERS Economic Research Service of the U.S. Department of Agriculture 

USDA-NASS National Agricultural Statistics Service of the U.S. Department of Agriculture 

WSSA Weed Science Society of America 

*Abbreviations of units of measurement and of physical and chemical quantities are used according to the 

standard format described in Instructions to Authors in the Journal of Biological Chemistry 

(http://www.jbc.org/). 
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73496 Canola 

I. Rationale for the Development of 73496 Canola 

I-A. Basis for the Request for a Determination of Nonregulated Status under 7 CFR §340.6 

The Animal and Plant Health Inspection Service (APHIS) of the U.S. Department of Agriculture (USDA) 

has responsibility, under the Plant Protection Act (7 U.S.C. 7701-7772) and the Plant Quarantine Act (7 

U.S.C. 151-167), to prevent the introduction or dissemination of plant pests into or within the United 

States. Part 340 regulates introduction of organisms altered or produced through genetic engineering 

which are plant pests or for which there is a reason to believe are plant pests. The APHIS regulations at 7 

CFR §430.6 provide that an applicant may petition APHIS to evaluate submitted data on the genetically 

engineered crop to determine that a regulated article does not present a plant pest risk and therefore 

should no longer be regulated. 

Pioneer Hi-Bred International, Inc. is submitting data for genetically engineered herbicide tolerant 73496 

canola and requests a determination from USDA - Animal and Plant Health Inspection Service (APHIS) 

that event DP-Ø73496-4 and any crosses with other nonregulated Brassica napus no longer be 

considered regulated articles under 7 CFR §340. 

I-B. Benefits of 73496 Canola 

Herbicide-tolerant canola varieties have provided growers with a very useful tool to help manage their 

canola crops. Herbicide-tolerant canola varieties have helped to increase the adoption of low- and no-till 

practices which are important in the minimization of soil erosion and also have helped to reduce the 

volume of herbicides applied to canola crops to control weeds (Brookes and Barfoot, 2010; Duke and 

Powles, 2009; Smyth et al., 2010). Additionally, the yield potential of canola can be optimized by the 

application of herbicides that remove weeds competing for soil nutrients and moisture. 

In particular, glyphosate has become a popular post-emergence herbicide for canola to control weeds 

that reduce yields. As a broad-spectrum herbicide, it is effective in controlling both grasses and annual 

broadleaf weeds, while possessing an excellent environmental profile and low mammalian toxicity. 

73496 canola was developed with glyphosate-tolerance to provide an alternative to existing herbicidetolerant canola products on the market. The commercialization of 73496 canola is not expected to have 

an impact on existing weed control practices and could replace the use of other glyphosate-tolerant 

canola products already commercialized. The availability of 73496 canola will provide growers with an 

additional market choice for herbicide-tolerant canola. 

I-C. Submissions to Other Regulatory Agencies 

A New Protein Consultation for the GAT4621 protein was submitted to FDA on January 31, 2007 and 

completed on October 7, 2009. A safety and nutritional assessment for feed and food derived from 

73496 canola will be submitted to FDA in the first quarter of 2011. 

Submission of a tolerance petition and supporting residue data to the U.S. Environmental Protection 

Agency (EPA) to amend the glyphosate tolerance to include N-acetylglyphosate for canola was submitted 

on February 18, 2011. 

Submissions for food, feed, and environmental approval were made to Canada in August 2010. 

Submissions will also be made in Mexico, Japan, South Korea, China, EU, and other countries as 

appropriate. Pioneer is committed to robust product stewardship prior to and continuing after all relevant 

authorizations are granted. A full commercial launch of 73496 canola will not occur until import regulatory 

approvals have been obtained in key canola import markets with functioning regulatory systems. 
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73496 Canola 

II. The Biology of Canola 

II-A. Canola as a Crop 

Biology documents on the unmodified plant species, canola (Brassica napus L.), have been published by 

the Canadian Food Inspection Agency (CFIA, 1994) and by the Organization for Economic Co-operation 

and Development (OECD, 1997). These documents provide background on the biology of B. napus

including: 

 information on use of canola as a crop plant 

 taxonomic status of Brassica

 identification methods 

 reproductive biology 

 centers of origin and diversity 

 crosses, including intra- and inter-specific/genus crosses and gene flow 

 agro-ecology, including information about cultivation, volunteers and weediness, soil ecology, and 

canola-insect interactions 

II-B. Description of the Non-Transformed Recipient Canola Line 

A doubled-haploid form of the Pioneer proprietary line, 1822B, was used as the recipient line for the 

production of 73496 canola. Line 1822B was chosen because it is receptive to transformation and is also 

an elite line (i.e., Pioneer proprietary line used for commercial products). The use of this doubled-haploid 

line ensured that the recipient line was homozygous at all genetic loci, which aided in the breeding and 

testing of the transformed events. 

III. Method of Development of 73496 Canola 

III-A. Description of Transformation, Selection, and Breeding 

A gel-purified DNA fragment isolated from plasmid PHP28181 (containing the gat4621 gene cassette) 

was used to generate 73496 canola. Refer to Section 3.1 for a detailed description of the gel-purified 

Hind III/Not I DNA fragment, PHP28181A, used for transformation. 

Microspores were prepared from donor line 1822B and transformed essentially as described (Chen and 

Tulsieram, 2007). Gold particles coated with the PHP28181A DNA fragment were used for biolistic 

transformation carried out as described (Klein et al., 1987). Transformed embryogenic microspores were 

cultured in fresh medium in dark conditions for 10 to 12 days, then under dim light for one to three weeks. 

Green embryos were transferred to fresh medium and cultured for two weeks at 4°C and then for four 

weeks at 25°C in the presence of glyphosate (0.1 mM) to select for glyphosate-tolerant transformants. 

Germinated shoots or plants were transferred to growth medium supplemented with glyphosate for 

selection. 

Plants that were regenerated from transformation and tissue culture (designated T0 plants) were selected 

for further characterization by molecular analyses, herbicide efficacy, and agronomic evaluations. Refer 

to Figure 1 for a schematic overview of the transformation and event development process for 73496 

canola. 

The subsequent breeding of 73496 canola proceeded as indicated in Figure 2 to produce specific 

generations for the characterization and assessments conducted. Table 1 indicates the breeding 

generations used for each of the analyses described in this submission. During the breeding and product 

development of 73496 canola, seed was produced for multiple purposes, including product development, 

research testing, and regulatory testing. Because there was not enough seed from any one generation 

for all of the regulatory studies listed in Table 1, seed was sourced from multiple generations of 73496 

canola. Where applicable, studies contained the appropriate near-isoline and/or commercial comparator. 

Each generation is equally representative of 73496 canola. 
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73496 Canola 

Assembly and isolation of DNA fragment PHP28181A containing gat4621 gene cassette 

↓

Transformation of microspores from 

1822B line using particle bombardment 

↓

Selection of transformation events based on tolerance to glyphosate 

Regeneration of T0 canola plants 

↓

Molecular characterization and evaluation of T0 plants 

↓

Selfing and molecular analysis to select homozygous plants 

↓

Nursery evaluation of agronomic performance and herbicide efficacy of subsequent generations 

↓

Multiple location herbicide efficacy and agronomic trials 

↓

Selection of event DP-Ø73496-4 as the commercial candidate 

↓

Continued field and laboratory studies to support product registration 

Continued breeding and testing of 73496 canola for product development 

Figure 1. Schematic Diagram of the Development of 73496 Canola 

T0 1822B

Self

T1

Self

T2

Self

T3

T4

X 1822R 

T3F1

Self

T3F2

T3F3

T3F4

X 5536F

F1*4

F1*5

Self

X 5300B

F1 *1

Self

BC1F1*1

BC2F1*1

BC3F1*1

X 5300B

X 5300B

X 5300B

BC2F1*3

X 6393B

X 6393B

X 6393B

BC1F1*3

X 6393B

BC3F1*3

X 6395B

X 5536F

Self

F1 F1 *2 *3

Self

T5

Figure 2. Breeding Diagram for 73496 Canola and Generations Used for Analyses
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73496 Canola 

Table 1. Generations and Comparators Used for Analysis of 73496 Canola

Analysis Generation Comparators and Commercial 

Reference Lines 

Genetic Characterization 

(Section V) T2, T3, F1*2, T3F2, T3F3 1822B, 1822R, and 6395B 

Inheritance 

(Section V-E) 

F1*1, *3, BC1F1*1, *3, BC2F1*1, *3, 

BC3F1*1, *3, T3F2 Not applicable 

Concentrations of GAT4621 

(Section V-B) F1*4 5536F x 1822R (near isoline) 

Compositional Assessment 

(Section VIII) F1*4 5536F x 1822R (near isoline) and 

Pioneer commercial lines 46A65, 

45H72, 45H73, 46H02, and 44A89 

Germination and Dormancy 

(Agronomic Performance) 

(Section VII-A) 

F1*5 5536F x 1822R (near isoline) and 

Pioneer commercial lines 45H72 

and 45H73 

Field Trial Evaluations (Agronomic 

Performance) 

(Section VII-B) 

T3 (7 locations) 

T5 (3 locations) 

1822B 

1822B 

III-B. Selection of Comparators for 73496 Canola 

To ensure an accurate evaluation of the effect of the transgene, a proper selection of comparator plants is 

important for the analysis of a transgenic plant. Control canola lines were used as comparators for 73496 

canola (Table 1). For all analyses, the near-isoline control plants had a genetic background similar to that 

of the 73496 canola generation used (i.e., >95% genetic similarity), but did not go through the 

transformation process. 

In addition, non-transgenic commercial canola lines (Table 1) were used to obtain tolerance intervals for 

the nutrient compositional assessment. These Pioneer commercial canola products were chosen to 

represent the full range of non-genetically modified varieties that would normally be planted commercially. 

The tolerance intervals were used to establish the range of natural variation of current commercial canola 

products with respect to key nutrients and anti-nutrients, and to determine the possible biological 

significance of any observed statistical differences between 73496 canola and commercial canola lines. 

IV. Donor Genes and Regulatory Sequences 

IV-A. Fragment PHP28181A Used in Transformation 

73496 canola was produced by biolistic transformation with the Hind III/Not I fragment, PHP28181A, from 

plasmid PHP28181. The PHP28181A fragment containing the UBQ10 promoter, the gat4621 gene, and 

the pinII terminator is shown in Figure 3, and the entire PHP28181 plasmid is shown in Figure 4. 
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73496 Canola 

PHP28181A

2112 bp

gat4621

UBQ10 Promoter pinII Terminator

Figure 3. Schematic Diagram of Fragment PHP28181A 

Schematic diagram of transformation fragment PHP28181A with the gat4621 gene and its regulatory elements 

indicated. The size of the fragment is 2112 base pairs (bp). 

PHP28181

4770 bp

bla (ApR)

gat4621

UBQ10 Promoter

pinII Terminator

Not I (2113)

Hind III (1)

colE1 ori

Figure 4. Schematic Diagram of Plasmid PHP28181

Schematic diagram of plasmid PHP28181 indicating the genetic elements. Hind III and Not I restriction enzyme 

sites flank the transformation fragment PHP28181A. The size of the plasmid is 4770 bp. 
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73496 Canola 

IV-B. Identity and Source of the gat4621 Gene Cassette on Fragment PHP28181A 

 

The genetic elements present in the transformation fragment PHP28181A are listed in Table 2. The 

gat4621 gene was optimized by a gene shuffling process of glyphosate acetyltransferase genes from 

Bacillus licheniformis. The GAT4621 protein, encoded by the gat4621 gene, confers tolerance to 

glyphosate-containing herbicides by acetylating glyphosate, thereby rendering it non-phytotoxic (Castle et 

al., 2004). 

B. licheniformis, the source organism for the gat4621 gene, is used for the production of a number of 

enzymes such as proteases and amylases that have wide application in the detergent industry. B. 

licheniformis has been used in the United States, Canada, and Europe in the fermentation industry for 

production of food enzymes (e.g., alpha-amylase, cyclodextrin glycosyltransferase, hemicellulase, 

proteases, and pullulanase; Rey et al., 2004). All B. licheniformis cultures available from the American 

Type Culture Collection (ATCC) are classified as Biosafety Level 1. Items that the ATCC classifies at 

Biosafety Level 1 have no known history of causing disease in humans or animals based on their 

assessment of potential risk using U.S. Public Health Service guidelines, with assistance provided by 

ATCC scientific advisory committees. 

The expression of the gat4621 gene is controlled by the Arabidopsis thaliana polyubiquitin promoter 

(UBQ10) (Norris et al., 1993). The terminator for the gat4621 gene is the 3’ terminator sequence from the 

proteinase inhibitor II gene of Solanum tuberosum (pinII terminator) (An et al., 1989; Keil et al., 1986). 

Table 2. Description of the Genetic Elements in Fragment PHP28181A 

Location on 

Fragment 

 (bp position) 

Genetic 

Element 

Size 

(base pairs) Description 

1 to 7 Polylinker 

Region 7 Region required for cloning genetic elements 

8 to 1312 UBQ10

Promoter 1305 

Version of the promoter region from Arabidopsis thaliana 

UBQ10 polyubiquitin gene (Norris et al., 1993) developed 

by E. I. duPont de Nemours and Company 

1313 to 1335 Polylinker 

Region 23 Region required for cloning genetic elements 

1336 to 1779 gat4621 Gene 444 Synthetic glyphosate N-acetyltransferase gene (Castle et 

al., 2004; Siehl et al., 2007) 

1780 to 1796 Polylinker 

Region 17 Region required for cloning genetic elements 

1797 to 2106 pinII 

Terminator 310 Terminator region from Solanum tuberosum proteinase inhibitor II gene (An et al., 1989; Keil et al., 1986) 

2107 to 2112 Polylinker 

Region 6 Region required for cloning genetic elements 
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73496 Canola 

V. Genetic Characterization of 73496 Canola 

V-A. Molecular Analysis Overview 

Molecular characterization of the inserted DNA evaluates the integrity of the introduced cassette and 

provides a confirmation that the elements of the expression cassette are intact. Genetic stability is 

evaluated to confirm the inheritance of the insertion and confirms the stability of the introduced trait 

through traditional breeding methods. 

The inserted DNA in 73496 canola was characterized by Southern blot analysis to evaluate the integrity 

and stability of the inserted gat4621 cassette. As described earlier in Section III-A, 73496 canola was 

produced by biolistic transformation with fragment PHP28181A. Fragment PHP28181A contains the 

gat4621 cassette containing the UBQ10 promoter, gat4621 gene, and pinII terminator. All probes used 

for the analysis are indicated on the schematic maps of PHP28181A and PHP28181 (Figures 5 and 6, 

respectively) and outlined in Table 3. Plasmid PHP28181 was used as a positive control for probe 

hybridization and to verify fragment sizes internal to PHP28181A. Individual plants of the T2, T3, T3F2, 

T3F3, and F1*2

 generations (refer to Figure 2 for the breeding diagram) were analyzed to determine the 

copy number of each of the genetic elements inserted into 73496 canola and to verify that the integrity of 

the PHP28181A fragment was maintained upon integration. 

The analyses confirmed that a single, intact PHP28181A DNA fragment was inserted into the genome of 

73496 canola (Section V-B1 and B2). In addition, these analyses also verified that the inserted DNA 

remained intact and stably integrated in 73496 canola. All five generations demonstrated identical 

hybridization patterns. These results confirmed the stability of the inserted DNA in 73496 canola across 

these five breeding generations. Based on these analyses, schematic maps of the inserted DNA in 

73496 canola were determined and are provided in Figure 7. 

All five generations were also analyzed to confirm the absence of plasmid backbone sequence, i.e. the 

plasmid region outside of isolated fragment PHP28181A. The results verified the absence of these 

backbone sequences in both 73496 canola (Section V-B3). 

To confirm inheritance of the insertion through traditional breeding, segregation analysis using genotypic 

and phenotypic assays were conducted (Section V-C). These results indicated that the inserted DNA 

and the herbicide-tolerance phenotype in 73496 canola segregate according to Mendel’s laws of 

segregation and are consistent with the finding of a single locus of insertion of the gat4621 cassette. The 

stability of the insertion and of the herbicide-tolerance phenotype was demonstrated in these generations 

of self- and cross-pollinations. 

Materials and methods for the molecular and genetic characterization of 73496 canola are described in 

Appendix 1. 

V-B. Results of Molecular Characterization of 73496 Canola by Southern Blot Analysis 

The integration pattern of the insertion in 73496 canola was investigated using Southern blot analysis with

Nco I digested genomic DNA to determine copy number and with Ssp I digested genomic DNA to 

determine insertion integrity. Copy number and integrity of each genetic element were determined in five 

generations of 73496 canola using probes specific to the UBQ10 promoter, gat4621 gene, and pinII 

terminator (Table 3, Figure 5). These analyses also verified that the inserted DNA remained intact and 

stably integrated. In addition, probes to the plasmid backbone region of PHP28181 located outside of the 

PHP28181A DNA fragment (Table 3, Figure 6) were used to show that these regions were not transferred 

to 73496 canola. 

Based on the Southern blot analyses described below, it was determined that a single, intact PHP28181A 

DNA fragment was inserted into the genome of 73496 canola as diagramed in the insertion map (Figure 

7) and that no region from the backbone of plasmid PHP28181 was inserted. In addition, these results 

confirmed the stability of the inserted DNA in 73496 canola across five breeding generations. 
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73496 Canola 

B1. Copy Number 

The Nco I digestion provides information about the number of copies of the PHP28181A DNA fragment 

that have been integrated into the genome of 73496 canola. The PHP28181A DNA fragment contains a 

single Nco I restriction enzyme site at bp position 1335 (Figure 5) and any additional sites would fall in the 

canola genome outside the fragment sequence. Therefore, hybridization with probes from the gat4621

cassette would indicate the number of copies of each element found in 73496 canola based on the 

number of hybridizing bands (e.g. one hybridizing band indicates one copy of the element). As the Nco I 

restriction site is located between the UBQ10 promoter and gat4621 gene, the promoter probe is 

expected to hybridize to a single band of greater than 1300 bp, and the gat4621 and pinII terminator 

probes are expected to hybridize to a single band of greater than 800 bp (Figure 5). Predicted and 

observed fragment sizes for 73496 canola with Nco I digestion are provided in Table 4. 

The UBQ10 promoter probe was hybridized to Nco I-digested genomic DNA from individual 73496 canola 

plants of the T2, T3, T3F2, T3F3, and F1*2

 generations (Table 4, Figure 8). A single fragment of greater 

than 8600 bp was detected in each 73496 canola plant sample (Table 4, Figure 8), indicating a single 

copy insertion of the promoter element. The gat4621 and pinII terminator probes were hybridized to the 

same Nco I-digested genomic DNA. Both probes hybridized to the same single band of approximately 

7000 bp in each 73496 canola plant sample (Table 4, Figures 9 and 10). 

The presence of single bands for each probe in this Southern blot analysis demonstrated the presence of 

a single copy of the PHP28181A DNA fragment in 73496 canola. The presence of identical hybridization 

patterns for each probe in all 73496 canola plants of the five generations analyzed demonstrated the 

stability of the DNA insertion during traditional breeding. 

B2. Insertion Integrity 

Ssp I digestion was used to verify that the inserted PHP28181A DNA fragment containing the gat4621

cassette was complete and intact in 73496 canola. Two Ssp I sites are present within the PHP28181A 

DNA fragment (bp positions 157 and 2043; Figure 5). Hybridization with the probes of the gat4621

cassette confirmed that all the elements were found on the expected internal 1886 bp fragment. 

Expected and observed fragment sizes for 73496 canola with Ssp I are provided in Table 4. All three 

probes are expected to hybridize to an internal fragment of 1886 bp (Table 4, Figure 5). Due to the 

locations of the Ssp I restriction sites within the UBQ10 promoter and pinII terminator elements (Figure 5), 

these two probes are also expected to hybridize to additional border fragments (Table 4). 

The UBQ10 promoter, gat4621, and pinII terminator probes all hybridized to a single insert-derived band 

of 1886 bp that matched the plasmid control band in each 73496 canola plant sample (Table 4, Figures 

11, 12, and 13). The UBQ10 promoter probe was expected to hybridize to a border band due to the Ssp I 

site within the UBQ10 promoter element. A faint border band of approximately 2200 bp was detected 

with the UBQ10 promoter probe (Table 4, Figure 11). A weakly hybridizing border band of approximately 

1100 bp was also detected with the pinII terminator probe (Table 4, Figure 13). The weak band resulted 

from a border fragment including the portion of the pinII terminator located 3’ to the Ssp I site at bp 2043 

of PHP28181A (Figure 5) and a second Ssp I site for this fragment located in the canola genome. These 

hybridizing border bands with the UBQ10 promoter and pinII terminator probes were observed in each 

73496 canola plant sample (Figures 11 and 13). 

The presence of the 1886 bp internal band with all probes and the border bands with the UBQ10

promoter and pinII terminator probes indicate that the PHP28181A fragment was inserted intact into 

73496 canola. Furthermore, the same hybridization pattern with each probe in the five generations 

confirmed the stability of the insertion in 73496 canola during traditional breeding. 

B3. Plasmid Backbone DNA Analysis 

Five generations of 73496 canola were analyzed by Southern blot analysis for the presence of DNA 

sequences from the PHP28181 plasmid backbone. Probes for sequences from the PHP28181 plasmid 
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73496 Canola 

backbone located outside the PHP28181A DNA fragment were used to determine if any plasmid 

backbone was inserted in 73496 canola during transformation. Three probes covered the entire plasmid 

backbone region and were used in a single hybridization solution. These probes would confirm if any 

region of the backbone outside the PHP28181A DNA fragment was transferred into 73496 canola. 

Genomic DNA from the T2, T3, T3F2, T3F3, and F1*2

 generations were digested with Nco I and 

hybridized to the backbone probes described above. No bands were observed in the 73496 canola or 

control canola samples, while the expected band of 4770 bp was seen in the plasmid positive control 

lanes (Figure 14). This confirms that no sequence from the PHP28181 plasmid backbone was inserted 

into 73496 canola during transformation. 

B4. Physical Map of the Inserted DNA in 73496 Canola 

Based on the Southern blot analysis, it was determined that a single, intact PHP28181A DNA fragment 

was inserted into the genome of 73496 canola. Using these data, a physical map of the inserted DNA in 

73496 canola showing the applicable restriction enzymes was developed and is provided in Figure 7. 

B5. Summary and Conclusions 

Southern blot analysis was conducted to characterize the DNA insertion in 73496 canola. The analysis 

confirmed that a single, intact PHP28181A DNA fragment had been inserted into the canola genome to 

produce 73496 canola. A single copy of each of the UBQ10 promoter, gat4621 gene, and pinII terminator 

genetic elements was present and the integrity of the PHP28181A DNA fragment was maintained upon 

integration. Identical hybridization patterns were observed across the T2, T3, T3F2, T3F3, and F1*2

generations, thus demonstrating stability of inheritance of the insertion in 73496 canola during traditional 

breeding procedures. In addition, Southern blot analysis verified the absence of plasmid backbone 

sequences in 73496 canola.
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73496 Canola 

Table 3. Description of DNA Probes Used for Southern Blot Hybridization 

Probe Name Genetic Element Figure 

Probe 

Position on 

PHP28181A 

DNA Fragment 

(bp to bp) 

Position on 

PHP28181 

Plasmid 

(bp to bp) 

Probe 

Length 

(bp) 

UBQ10

promotera UBQ10 promoter Figure 5 

probe A 

26 to 699 

720 to 1286 

26 to 699 

720 to 1286 

674 

567 

gat4621 gat4621 gene Figure 5 

probe B 1336 to 1770 1336 to 1770 435 

pinII terminator pinII terminator Figure 5 

probe C 1849 to 2082 1849 to 2082 234 

Backbone 1 Plasmid between Not I 

and bla

Figure 6 

probe 1 N/A 2120 to 2762 643 

bla (ApR) Ampicillin resistance 

gene 

Figure 6 

probe 2 N/A 2736 to 3566 831 

Backbone 2 Plasmid between 

bla and Hind III 

Figure 6 

probe 3 N/A 3503 to 4770 1268 

N/A-Not Applicable, these are not present on the PHP28181A DNA fragment. 

a

 Two non-overlapping segments were generated for this probe and were combined for hybridization. The bp 

positions provided are the positions of each different segment. 
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73496 Canola 

Table 4. Summary of Expected and Observed Hybridization Fragments on Southern Blots for 

73496 Canola 

Probe Enzyme 

Digestion 

Figure 

Expected 

Fragment 

Size in 

73496 Canola 

(bp)a

Expected 

Fragment Size in 

Plasmid 

PHP28181 (bp)b

Observed 

Fragment Size in 

73496 Canola 

(bp)c

UBQ10 Promoter 

Nco I 8 >1300 (border) 4770 >8600 

Ssp I 11 >200 (border) 

1886 

2208 

1886 

~2200 

1886d

gat4621 Nco I 9 >800 (border) 4770 ~7000 

Ssp I 12 1886 1886 1886d

pinII Terminator 

Nco I 10 >800 (border) 4770 ~7000 

Ssp I 13 1886 

>100 (border) 

1886 

676 

1886d

~1100 

Plasmid Backbonee Nco I 14 none 4770 none 

a

 Size based on map of fragment PHP28181A in Figure 5. Border fragment size rounded to nearest 100 bp. b

 Size based on plasmid map of PHP28181 in Figure 6. 

c

 Observed fragment sizes are approximated from the digoxigenin (DIG)-labeled DNA Molecular Weight Marker 

VII fragments on the Southern blots. Due to the inability to determine exact sizes on the blot, all approximated 

values are rounded to the nearest 100 bp. 

d

 Size is same as expected because of equivalent migration with plasmid positive control. e

 The plasmid backbone probe includes three fragments covering the entire plasmid backbone sequence in a 

single hybridization solution. 

gat4621

UBQ10 Promoter pinII Terminator Nco I (1335)

Ssp I (157) Ssp I (2043)

A B C

Figure 5. Restriction Enzyme Map of PHP28181A DNA Fragment 

Schematic map of fragment PHP28181A from plasmid PHP28181 used for transformation of canola. Location of

genetic elements and Ssp I and Nco I restriction enzyme sites are indicated. Fragment size is 2112 bp. The 

locations of the probes used are shown as boxes in the lower part of the map and are identified as follows: A = 

UBQ10 promoter probe, B = gat4621 probe, and C = pinII terminator probe. 
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73496 Canola 

Figure 6. Restriction Enzyme Map of Plasmid PHP28181 

Schematic map of plasmid PHP28181 indicating the location of genetic elements and Not I, Ssp I, Nco I, and 

Hind III restriction enzyme sites. Hind III and Not I sites flank the fragment PHP28181A (Figure 5) used for 

transformation of canola. The black arcs inside the plasmid map indicate the locations of the three probes used 

to confirm the absence of plasmid backbone sequences and are identified as follows: 1 = Backbone 1 probe, 2 = 

bla (ApR) probe, and 3 = Backbone 2 probe.

PHP28181A

Nco I >8600 bp ~7000 bp

Ssp I ~2200 bp 1886 bp ~1100 bp

Nco I Ssp I Nco I gat4621

UBQ10 Promoter Nco pinII Terminator I 

Ssp I Ssp I Ssp I

Figure 7. Physical Map of the Insertion in 73496 Canola 

Schematic map of the insertion in 73496 canola based on Southern blot analysis. The flanking canola genome is 

represented by the horizontal dotted line. A single, intact copy of the PHP28181A DNA fragment integrated into 

the canola genome. Nco I and Ssp I restriction enzyme sites are indicated with the sizes of observed fragments 

on Southern blots shown below the map in bp. The locations of restriction enzyme sites outside the PHP28181A 

DNA fragment are not shown to scale. 
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73496 Canola 

Figure 8. Southern Blot Analysis of 73496 Canola: Nco I Digestion and UBQ10 Promoter Probe 

Genomic DNA isolated from leaf tissue from individual plants from five generations of 73496 canola and control 

canola lines was digested with Nco I and hybridized to the UBQ10 promoter probe. Approximately 4 μg of 

genomic DNA was digested and loaded per lane. Positive control lanes include plasmid PHP28181 and 4 μg of 

control canola DNA. Sizes of the DIG VII molecular weight markers are indicated adjacent to the blot image in 

kilobases (kb). 

Lane Sample Lane Sample 

1 3 copy PHP28181 + 1822R control 11 73496 canola / plant 8 (T3 generation) 

2 1 copy PHP28181 + 1822R control 12 73496 canola / plant 9 (T3F2 generation)

3 DIG VII molecular weight marker 13 73496 canola / plant 10 (T3F2 generation)

4 1822B control 14 73496 canola / plant 14 (T3F3 generation)

5 1822R control 15 73496 canola / plant 15 (T3F3 generation)

6 6395B control 16 73496 canola / plant 19 (F1*2 generation)

7 Blank 17 73496 canola / plant 20 (F1*2 generation)

8 73496 canola / plant 1 (T2 generation) 18 DIG VII molecular weight marker 

9 73496 canola / plant 3 (T2 generation) 19 1 copy PHP28181 + 6395B control 

10 73496 canola / plant 7 (T3 generation) 20 3 copy PHP28181 + 6395B control 

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

kb

8.6

7.4

6.1

4.9

3.6

2.8

1.95

1.88

1.51

1.48

1.2

0.99

0.72

kb

8.6

7.4

6.1

4.9

3.6

2.8

1.95

1.88

1.51

1.48

1.2

0.99

0.72
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73496 Canola 

Figure 9. Southern Blot Analysis of 73496 Canola: Nco I Digestion and gat4621 Probe 

Genomic DNA isolated from leaf tissue from individual plants from five generations of 73496 canola and control 

canola lines was digested with Nco I and hybridized to the gat4621 probe. Approximately 4 μg of genomic DNA 

was digested and loaded per lane. Positive control lanes include plasmid PHP28181 and 4 μg of control canola 

DNA. Sizes of the DIG VII molecular weight markers are indicated adjacent to the blot image in kilobases (kb). 

Lane Sample Lane Sample 

1 3 copy PHP28181 + 1822R control 11 73496 canola / plant 8 (T3 generation) 

2 1 copy PHP28181 + 1822R control 12 73496 canola / plant 9 (T3F2 generation)

3 DIG VII molecular weight marker 13 73496 canola / plant 10 (T3F2 generation)

4 1822B control 14 73496 canola / plant 14 (T3F3 generation)

5 1822R control 15 73496 canola / plant 15 (T3F3 generation)

6 6395B control 16 73496 canola / plant 19 (F1*2 generation)

7 Blank 17 73496 canola / plant 20 (F1*2 generation)

8 73496 canola / plant 1 (T2 generation) 18 DIG VII molecular weight marker 

9 73496 canola / plant 3 (T2 generation) 19 1 copy PHP28181 + 6395B control 

10 73496 canola / plant 7 (T3 generation) 20 3 copy PHP28181 + 6395B control 

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

kb

8.6

7.4

6.1

4.9

3.6

2.8

1.95

1.88

1.51

1.48

1.2

0.99

0.72

kb

8.6

7.4

6.1

4.9

3.6

2.8

1.95

1.88

1.51

1.48

1.2

0.99

0.72
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73496 Canola 

Figure 10. Southern Blot Analysis of 73496 Canola: Nco I Digestion and pinII Terminator Probe 

Genomic DNA isolated from leaf tissue from individual plants from five generations of 73496 canola and control 

canola lines was digested with Nco I and hybridized to the pinII terminator probe. Approximately 4 μg of genomic 

DNA was digested and loaded per lane. Positive control lanes include plasmid PHP28181 and 4 μg of control 

canola DNA. Sizes of the DIG VII molecular weight markers are indicated adjacent to the blot image in kilobases 

(kb). 

Lane Sample Lane Sample 

1 3 copy PHP28181 + 1822R control 11 73496 canola / plant 8 (T3 generation) 

2 1 copy PHP28181 + 1822R control 12 73496 canola / plant 9 (T3F2 generation)

3 DIG VII molecular weight marker 13 73496 canola / plant 10 (T3F2 generation)

4 1822B control 14 73496 canola / plant 14 (T3F3 generation)

5 1822R control 15 73496 canola / plant 15 (T3F3 generation)

6 6395B control 16 73496 canola / plant 19 (F1*2 generation)

7 Blank 17 73496 canola / plant 20 (F1*2 generation)

8 73496 canola / plant 1 (T2 generation) 18 DIG VII molecular weight marker 

9 73496 canola / plant 3 (T2 generation) 19 1 copy PHP28181 + 6395B control 

10 73496 canola / plant 7 (T3 generation) 20 3 copy PHP28181 + 6395B control 

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

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Figure 11. Southern Blot Analysis of 73496 Canola: Ssp I Digestion and UBQ10 Promoter Probe 

Genomic DNA isolated from leaf tissue from individual plants from five generations of 73496 canola and control 

canola lines was digested with Ssp I and hybridized to the UBQ10 promoter probe. Approximately 4 μg of 

genomic DNA was digested and loaded per lane. Positive control lanes include plasmid PHP28181 and 4 μg of 

control canola DNA. Sizes of the DIG VII molecular weight markers are indicated adjacent to the blot image in 

kilobases (kb). 

Note: A faint band is visible on the X-ray film at about 2.2 kb in lanes 8 through 17 (may not appear on printed 

copy). 

Lane Sample Lane Sample 

1 3 copy PHP28181 + 1822R control 11 73496 canola / plant 8 (T3 generation) 

2 1 copy PHP28181 + 1822R control 12 73496 canola / plant 9 (T3F2 generation)

3 DIG VII molecular weight marker 13 73496 canola / plant 10 (T3F2 generation)

4 Blank 14 73496 canola / plant 14 (T3F3 generation)

5 1822B control 15 73496 canola / plant 15 (T3F3 generation)

6 1822R control 16 73496 canola / plant 19 (F1*2 generation)

7 6395B control 17 73496 canola / plant 20 (F1*2 generation)

8 73496 canola / plant 1 (T2 generation) 18 DIG VII molecular weight marker 

9 73496 canola / plant 3 (T2 generation) 19 1 copy PHP28181 + 6395B control 

10 73496 canola / plant 7 (T3 generation) 20 3 copy PHP28181 + 6395B control 
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73496 Canola 

Figure 12. Southern Blot Analysis of 73496 Canola: Ssp I Digestion and gat4621 Probe 

Genomic DNA isolated from leaf tissue from individual plants from five generations of 73496 canola and control 

canola lines was digested with Ssp I and hybridized to the gat4621 probe. Approximately 4 μg of genomic DNA 

was digested and loaded per lane. Positive control lanes include plasmid PHP28181 and 4 μg of control canola 

DNA. Sizes of the DIG VII molecular weight markers are indicated adjacent to the blot image in kilobases (kb). 

Lane Sample Lane Sample 

1 3 copy PHP28181 + 1822R control 11 73496 canola / plant 8 (T3 generation) 

2 1 copy PHP28181 + 1822R control 12 73496 canola / plant 9 (T3F2 generation)

3 DIG VII molecular weight marker 13 73496 canola / plant 10 (T3F2 generation)

4 Blank 14 73496 canola / plant 14 (T3F3 generation)

5 1822B control 15 73496 canola / plant 15 (T3F3 generation)

6 1822R control 16 73496 canola / plant 19 (F1*2 generation)

7 6395B control 17 73496 canola / plant 20 (F1*2 generation)

8 73496 canola / plant 1 (T2 generation) 18 DIG VII molecular weight marker 

9 73496 canola / plant 3 (T2 generation) 19 1 copy PHP28181 + 6395B control 

10 73496 canola / plant 7 (T3 generation) 20 3 copy PHP28181 + 6395B control 

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Figure 13. Southern Blot Analysis of 73496 Canola: Ssp I Digestion and pinII Terminator Probe 

Genomic DNA isolated from leaf tissue from individual plants from five generations of 73496 canola and control 

canola lines was digested with Ssp I and hybridized to the pinII terminator probe. Approximately 4 μg of genomic 

DNA was digested and loaded per lane. Positive control lanes include plasmid PHP28181 and 4 μg of control 

canola DNA. Sizes of the DIG VII molecular weight markers are indicated adjacent to the blot image in kilobases 

(kb). 

Note: A faint band is visible on the x-ray film at about 1.1 kb in lanes 8 through 17 (may not appear on printed 

copy).

Lane Sample Lane Sample 

1 3 copy PHP28181 + 1822R control 11 73496 canola / plant 8 (T3 generation) 

2 1 copy PHP28181 + 1822R control 12 73496 canola / plant 9 (T3F2 generation)

3 DIG VII molecular weight marker 13 73496 canola / plant 10 (T3F2 generation)

4 Blank 14 73496 canola / plant 14 (T3F3 generation)

5 1822B control 15 73496 canola / plant 15 (T3F3 generation)

6 1822R control 16 73496 canola / plant 19 (F1*2 generation)

7 6395B control 17 73496 canola / plant 20 (F1*2 generation)

8 73496 canola / plant 1 (T2 generation) 18 DIG VII molecular weight marker 

9 73496 canola / plant 3 (T2 generation) 19 1 copy PHP28181 + 6395B control 

10 73496 canola / plant 7 (T3 generation) 20 3 copy PHP28181 + 6395B control 

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

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Figure 14. Southern Blot Analysis of 73496 Canola: Nco I Digestion and Plasmid Backbone Probe 

Genomic DNA isolated from leaf tissue from individual plants from five generations of 73496 canola and control 

canola lines was digested with Nco I and hybridized to the combined plasmid backbone probe. Approximately 4 

μg of genomic DNA was digested and loaded per lane. Positive control lanes include plasmid PHP28181 and 4 

μg of control canola DNA. Sizes of the DIG VII molecular weight markers are indicated adjacent to the blot 

image in kilobases (kb). 

Note: Extra bands in Lane 3 near and below the 0.72 kb band are due to hybridization of the probe to the ΦX174 

RF DNA/Hae III fragments used to monitor gel electrophoresis progress.

Lane Sample Lane Sample 

1 3 copy PHP28181 + 1822R control 11 73496 canola / plant 8 (T3 generation) 

2 1 copy PHP28181 + 1822R control 12 73496 canola / plant 9 (T3F2 generation)

3 DIG VII molecular weight marker 13 73496 canola / plant 10 (T3F2 generation)

4 1822B control 14 73496 canola / plant 14 (T3F3 generation)

5 1822R control 15 73496 canola / plant 15 (T3F3 generation)

6 6395B control 16 73496 canola / plant 19 (F1*2 generation)

7 Blank 17 73496 canola / plant 20 (F1*2 generation)

8 73496 canola / plant 1 (T2 generation) 18 DIG VII molecular weight marker 

9 73496 canola / plant 3 (T2 generation) 19 1 copy PHP28181 + 6395B control 

10 73496 canola / plant 7 (T3 generation) 20 3 copy PHP28181 + 6395B control 

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

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V-C. Inheritance and Genetic Stability of the Introduced Trait in 73496 Canola 

Inheritance of the inserted DNA and the herbicide-tolerance phenotype in 73496 canola was evaluated to 

ensure stability of the trait during the plant breeding process and to confirm the trait was at a single 

genetic locus. Segregating generations of 73496 canola (T3F2, BC1F1*1,*3, BC2F1*1,*3, and BC3F1*1,*3) 

and one non-segregating generation (F1*1,*3) were evaluated. The breeding history of these five 

generations is shown in the breeding diagram (Section III-A, Figure 2). Those populations with the 

superscript “*1” or “*3” designation were populations from two different genetic backgrounds (Figure 2). 

Chi-square analysis was conducted on the four segregating generations to determine if the observed 

segregation ratios were consistent with the expected ratios. The assay methods and statistical analysis 

for the trait inheritance data are described in Appendix 1. 

The presence of the 73496 event insertion was determined by event-specific and gat4621 gene-specific 

endpoint PCR analyses performed on leaf punches from seedlings of each generation. The herbicide 

tolerance phenotype was determined by treating the plants with herbicide and by visually evaluating each 

plant for the presence of herbicide injury. A positive plant exhibited no herbicidal injury and a negative 

plant exhibited severe herbicide injury. 

Results from the segregation analysis are provided in Table 5. For one of the plants, PCR analyses were 

inconclusive and were unable to be repeated, and therefore were excluded from the statistical analyses. 

However, this excluded plant was evaluated for the herbicide-tolerance phenotype and this information 

was reported and analyzed. For those plants where both PCR and herbicide-tolerance phenotype data 

were obtained, all analyses correlated (e.g., plants that were positive by PCR analyses were also tolerant 

to the herbicide). To confirm that the inserted DNA and the herbicide-tolerance phenotype segregate 

according to Mendel’s laws of genetics, chi-square analysis was performed separately for the PCR and 

the herbicide-tolerance phenotype data. All P-values were greater than 0.05, with the exception of the 

BC1F1*1,*3 generation of 73496 canola, indicating that the observed segregation ratio was consistent with 

the expected ratio (Table 5). 

In the case of the BC1F1*1 and BC1F1*3 populations, the observed segregation ratios were not consistent 

with the expected 1:1 (positive: negative) ratio (Table 5) and this was attributed to the hand pollinations 

that were performed to generate the populations. The BC1F1*1 and BC1F1*3 populations were crosspollinated by hand with a control canola plant in order to generate a population segregating 1:1. This 

process required removal of pollen-producing anthers from several fertile flower buds of a hemizygous 

73496 canola plant. It is likely that some of the flower buds were self-pollinated inadvertently, generating 

a 3:1 segregating population, prior to the cross-pollination step and resulted in a pool of seed that was a 

mixture of 3:1 and 1:1 segregating populations. Each of the other 73496 canola generations that were 

analyzed were either derived from the same progenitor as the BC1F1 generation (T3F2), a parent of the 

BC1F1 generation (F1*1,*3), or derived directly from this BC1F1 generation (BC2F1*1,*3 and BC3F1*1,*3) 

(Figure 2). The fact that results from these other four generations were consistent with expectations 

indicate that the BC1F1*1 and BC1F1*3 populations were not representative of the segregation of the trait 

and that the insertion is segregating as expected. 

These results indicate that the inserted DNA and the herbicide-tolerance phenotype in 73496 canola 

segregate according to Mendel’s laws of segregation and were consistent with the finding of a single 

locus of insertion of the gat4621 cassette. The stability of the insertion and of the herbicide-tolerance 

phenotype was demonstrated in these generations of self- and cross-pollinations. 
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Table 5. Summary of Genotypic and Phenotypic Results for 73496 Canola 

73496 

Canola 

Generation 

Expected 

Segregation 

Ratio 

Seed 

Source Analysis 

Observed Values 

(n = 100a

) 

Statistical 

Analysis 

Positive Negative ChiSquareb Pvalue 

T3F2 3:1 T3F2 

PCR 69 30 1.4848 0.2230 

Herbicide tolerance assay 69 31 1.9200 0.1659 

F1 NonSegregating

F1 *1

PCR 

49 1 

NA

F1 c

*3 50 0 

F1 *1

Herbicide tolerance assay 

49 1 

F1 *3 50 0 

BC1F1 1:1 

BC1F1 *1

PCR 

32 18 3.9200 0.0477d

BC1F1 *3 32 18 3.9200 0.0477d

BC1F1 *1

Herbicide tolerance assay 

32 18 3.9200 0.0477d

BC1F1 *3 32 18 3.9200 0.0477d

BC2F1 1:1 

BC2F1 *1

PCR 

29 21 1.2800 0.2579 

BC2F1 *3 26 24 0.0800 0.7773 

BC2F1 *1

Herbicide tolerance assay 

29 21 1.2800 0.2579 

BC2F1 *3 26 24 0.0800 0.7773 

BC3F1 1:1 

BC3F1 *1

PCR 

24 26 0.0800 0.7773 

BC3F1 *3 27 23 0.3200 0.5716 

BC3F1 *1

Herbicide tolerance assay 24 26 0.0800 0.7773 

BC3F1 *3 27 23 0.3200 0.5716 a n = 99 for statistical analysis of T3F2 PCR results (PCR results for one T3F2 plant were inconclusive and were 

therefore excluded). 

b Degrees of freedom = 1 c Chi-square test is not applicable (NA) for testing a non-segregating population (i.e. a nominal proportion of 1.0). d Statistically significant difference, P-value <0.05 

V-D. Conclusions on Molecular Characterization and Genetic Stability of 73496 Canola 

Southern blot analysis was conducted to characterize the DNA insertion in 73496 canola. The analysis 

confirmed that a single, intact PHP28181A DNA fragment was inserted into the canola genome. A single 

copy of each of the UBQ10 promoter, gat4621 gene, and pinII terminator genetic elements was present 

and the integrity of the PHP28181A DNA fragment was maintained. Southern blot analysis on five 

generations confirmed the stability of inheritance of the DNA insertion during traditional breeding 

procedures. In addition, Southern blot analysis verified the absence of plasmid backbone sequences in 

73496 canola.

The inheritance and genetic stability of the inserted DNA in 73496 canola was examined in additional 

generations by event-specific and gene-specific PCR assays and by evaluation of herbicide-tolerance in 

the population. Four segregating generations and one non-segregating generation were evaluated for 

consistency with the expected segregation ratio of the population. The results of this analysis were 

consistent with the finding of a single locus of insertion of the gat4621 cassette in 73496 canola that 

segregated according to Mendelian rules of inheritance. The stability of the insertion and of the herbicidetolerance phenotype was demonstrated in these populations. 

Together, these analyses confirmed the stability of the inserted DNA and its corresponding herbicidetolerance trait during traditional breeding of 73496 canola. 
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73496 Canola 

VI. Characterization of the Introduced GAT4621 Protein 

73496 canola was modified to express the GAT4621 protein to confer glyphosate-tolerance. The identity, 

deduced amino acid sequence, enzymatic activity, derivation, protein characterization, and 

concentrations of the GAT4621 protein in 73496 canola tissues are described below. 

VI-A. The GAT4621 Protein 

A1. Identity of the GAT4621 Protein 

The GAT4621 protein is a variant of N-acetyltransferase protein sequences derived from Bacillus 

licheniformis, a gram positive saprophytic bacterium that is widespread in nature and thought to 

contribute substantially to nutrient cycling due to the diversity of enzymes produced by members of its 

species. GAT4621 is 147 amino acids in length and has a molecular weight of 16.5 kDa (Figure 15). 

Figure 15. Deduced Amino Acid Sequence of the GAT4621 Protein 

The introduced gat4621 coding sequence in 73496 canola is identical to the gat4621 coding sequence 

present in the previously approved maize event DP-Ø9814Ø-6 (FDA, 2008; Rood, 2007b; USDA, 2009). 

As discussed in Section VI-B, the GAT4621 protein produced in 73496 canola demonstrated 

physicochemical equivalence to the microbial-produced GAT4621 protein that was used in the protein 

safety studies that were conducted for maize event DP-Ø9814Ø-6. These bridging data also support the 

conclusion that the GAT4621 protein expressed in 73496 canola and in maize event DP-Ø9814Ø-6 are 

equivalent. 

A2. Characteristics and Enzymatic Activity of GAT Proteins 

Glyphosate acetyltransferase (GAT) proteins are members of the GCN 5-related family of 

N-acetyltransferases (also known as the GNAT superfamily). The GNAT superfamily is one of the largest 

enzyme superfamilies recognized to date with over 10,000 representatives from plants, animals and 

microbes. Members of the GNAT superfamily all contain highly conserved GNAT motifs but have high 

sequence diversity (Vetting et al., 2005). The GNAT proteins are known to have a number of metabolic 

functions including detoxification (Dyda et al., 2000). In particular, and as described below, GAT proteins 

can confer herbicide tolerance by detoxification of the broad-spectrum herbicide glyphosate. 

The enzyme 5-enolpyruvylshikimate-3-phosphate synthase (EPSPS; 3-phophoshikimate 1-carboxyvinyltransferase; EC2.5.1.19) (Steinrücken and Amrhein, 1980) is the sixth enzyme of the shikimic acid 

pathway, which is essential for the biosynthesis of aromatic amino acids (L-phenylalanine, L-tyrosine and 

L-tryptophan) and chorismate-derived secondary metabolites in algae, higher plants, bacteria, and fungi 

(Kishore and Shah, 1988). EPSPS has been identified as the primary target of glyphosate [N-

(phosphonomethyl) glycine], which is a nonselective, broad-spectrum, foliar-applied herbicide first 

commercialized in 1974 and widely used for the management of annual, perennial, and biennial 

herbaceous species of grasses, sedges, and broadleaf weeds, as well as woody brush and tree species 

(Baylis, 2000; Bradshaw et al., 1997). Mechanisms for conferring tolerance to glyphosate herbicide in 

genetically engineered plants have included the introduction of microbial variants of EPSPS that are 

insensitive to glyphosate (e.g., CP4 EPSPS from Agrobacterium tumefaciens) or mutated forms of 

endogenous plant EPSP synthases (e.g., modified EPSPS from Zea mays). 
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GAT proteins provide an alternative mechanism of tolerance to glyphosate by detoxifying glyphosate to 

the non-phytotoxic form, N-acetylglyphosate (Figure 16). This detoxification mechanism is similar to that 

of the phosphinothricin acetyltransferase (PAT or BAR) enzymes from Streptomyces, which detoxify 

glufosinate ammonium herbicides by adding an acetyl group (De Block et al., 1987). As shown in Figure 

16, GAT enzymes acetylate the secondary amine of glyphosate using acetyl coenzyme A as an acetyl 

donor (Castle et al., 2004). Transgenic expression of GAT proteins was shown to confer glyphosate 

tolerance in several plant species (Castle et al., 2004) and is also the basis for previously approved maize 

(DP-Ø9814Ø-6) (FDA, 2008; Rood, 2007b; USDA, 2009) and soybean (DP-356Ø43-5) events (FDA, 

2007; Rood, 2006; USDA, 2008). 

Figure 16. Enzymatic Activity of the GAT4621 Protein

A3. Derivation of the GAT4621 Protein 

The derivation of the GAT4621 enzyme was discussed in submissions to the USDA and FDA for maize 

event DP-Ø9814Ø-6 (FDA, 2008; Rood, 2007b; USDA, 2009) and was discussed in the New Protein 

Consultation that was provided to FDA (FDA, 2009; Rood, 2007a; Appendix 10). In summary, the 

gat4621 gene was produced using a DNA shuffling process employing three native gat genes from 

Bacillus licheniformis as parental templates (Castle et al., 2004). The gat4621 gene variant was the 

result of 11 rounds of gene shuffling and the encoded GAT4621 enzyme exhibited a 7000-fold increase in 

catalytic activity relative to the native enzymes, using glyphosate as a substrate (Siehl et al., 2005). The 

gat4621 gene was further optimized for plant expression by conservative codon substitutions and a codon 

addition to improve protein production in plants (a GCT codon for alanine was inserted at amino acid 

position 2).

The GAT4621 protein, encoded by the gat4621 gene, is 75–78% identical and 90–91% similar at the 

amino acid level to each of the three native GAT enzymes from which it was derived, compared to 94% 

identity of each of the native enzymes to each other (Table 6). Amino acid similarity is defined as the 

percentage of identical amino acids plus the percentage of “conservative” changes from one amino acid 

to another with the same functional side chain group (e.g., alanine to valine, or lysine to arginine) 

between two proteins. Therefore, the percent similarity is always equal to or greater than the percent 

identity between two proteins. There are 32 to 36 amino acid changes (22 to 23 of which are 

conservative) between the shuffled GAT4621 protein and any one of the original three native GAT 

proteins. 
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Table 6. Comparison of Amino Acid Sequence Similarity Between Parental GAT Enzymes and 

GAT4621 

Percent Identity or Similarity of Amino Acid Sequences 

GAT (strain 401) GAT (strain B6) GAT (strain DS3) GAT4621

GAT (strain 401) 100% 94% Identical 94% Identical 78% Identical 

91% Similar 

GAT (strain B6) 100% 94% Identical 76% Identical 

91% Similar 

GAT (strain DS3) 100% 75% Identical 

90% Similar 

GAT4621 100% 

VI-B. Physicochemical Characterization of the GAT4621 Protein 

The safety of GAT4621 protein was evaluated and was assessed previously by FDA as part of New 

Protein Consultation 005 (FDA, 2009; Rood, 2007a; Appendix 10). That assessment considered the 

allergenicity and toxicity of the GAT4621 protein, which used a microbial-expressed GAT4621 protein as 

a means to evaluate protein safety in certain studies. The microbial-expressed GAT4621 protein was 

characterized for maize event DP-Ø9814Ø-6 and was used to make relevant comparisons to the 

GAT4621 protein in 73496 canola to demonstrate equivalence and to confirm suitability of bridging to the 

previous GAT4621 safety assessments. 

In order to verify the equivalence of the in planta GAT4621 to the microbial-produced protein and confirm 

the microbial-produced protein was appropriate for the safety assessment studies, a physicochemical 

characterization of the GAT4621 protein in 73496 canola was conducted. This characterization of the in 

planta GAT4621 protein included the following analyses: 

 Molecular weight and immunochemical cross-reactivity by sodium dodecyl sulfate polyacrylamide 

gel electrophoresis (SDS-PAGE) and western immunoblot analysis 

 N-terminal amino acid sequencing to confirm protein identity 

 Mass determination and protein identity using tryptic peptide mapping by matrix assisted laser 

desportion ionization mass spectrometry (MALDI-MS) 

 Glycoprotein staining to determine potential post-translational modification (glycosylation) 

A detailed description of the methods used in the equivalency studies and the resulting data are included 

in Appendix 4. 

Utilizing the above analyses, the equivalency of GAT4621 protein expressed in E. coli to the protein 

expressed in 73496 canola was demonstrated. Therefore, the GAT4621 protein derived from the 

microbial expression system was appropriate for utilization in safety assessment studies as a proxy for 

the GAT4621 protein expressed in 73496 canola. Microbial GAT4621 protein was used for in vitro

digestion and acute toxicology safety assessment studies summarized in Section VI-D. 

VI-C. Concentrations of GAT4621 in 73496 Canola 

In order to understand the level expression of the GAT4621 protein in 73496 canola and the potential 

exposure, the concentration of the protein was measured in relevant plant tissues. Expression of the 

GAT4621 protein is driven by the constitutive UBQ10 promoter from Arabidopsis thaliana and would be 

expected to be measurable in all tissues tested from 73496 canola. 
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73496 Canola 

The range of expression of the GAT4621 protein in 73496 canola was determined by quantitative enzyme 

linked immunosorbent assay (ELISA) of samples of seed, roots, and whole plants (i.e., the entire 

above-ground portion of the plant) obtained from up to six field trial locations in Canada and the United 

States in 2009 (Appendix 3 for ELISA materials and methods). Near-isoline canola plants were grown 

and sampled as controls. 

As indicated below, tissue samples were collected at various developmental stages described by 

Lancashire et al. (1991): 

 BBCH15 – five true leaves unfolded (whole plant sample) 

 BBCH33 – three visibly extended internodes (whole plant sample) 

 BBCH65 – full flowering; 50% of flowers open on main raceme, older petals falling (whole plant 

and root samples) 

 BBCH90 – senescence (seed sample) 

As expected, quantifiable amounts of GAT4621 protein were detected in each plant tissue tested from 

73496 canola (Table 7). The range of mean concentrations in whole plant samples across growth stages 

was 5.2-6.9 ng/mg dry weight. The mean concentration of the GAT4621 protein measured in 73496 root 

was 6.6 ng/mg dry weight. The mean concentration in seed was 6.2 ng/mg dry weight. 

Table 7. Concentration of GAT4621 Protein in 73496 Canola Plant Tissues 

Tissue

GAT4621 Concentration [ng/mg dry weight ± SD (range)]a

Plant Growth Stageb

BBCH15 BBCH33 BBCH65 BBCH90 

Whole 

Plant 

n=24 

6.9 ± 1.3 

(3.9–10) 

5.3 ± 1.2 

(3.1–8.4) 

5.2 ± 0.88 

(3.9–7.6) NC 

Root 

n=24 NCc

 NC 6.6 ± 2.4 

(3.9–13) NC 

Seed 

n=20 NC NC NC 6.2 ± 0.94 (4.8–8.4) 

a

 Values are expressed as the mean of four replicate tissue samples collected from each of six locations except seed 

samples, which were only collected from five locations. SD = standard deviation; range, in parentheses, denotes the 

lowest and highest individual value across sites. b

 Plant growth stages: BBCH15 – five true leaves unfolded; BBCH33 – three visibly extended internodes; BBCH65 – 

full flowering; BBCH90 – senescence (Lancashire et al., 1991). c

 NC = Not collected.
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VI-D. Summary of the Food and Feed Safety Assessment for the GAT4621 Protein

A food and feed safety assessment was conducted to assess the allergenicity and toxicity potential of the 

GAT4621 protein. A detailed assessment of the food and feed safety of the GAT4621 protein was 

submitted to FDA on January 31, 2007 as part of New Protein Consultation (NPC) 005 and was 

completed on October 7, 2009 (FDA, 2009; Rood, 2007a). A summary of the information in NPC 005, 

along with recently updated bioinformatic analyses on GAT4621 are described below. Additional 

information on 73496 canola, including human and livestock exposure assessments, will be submitted to 

FDA as part of the consultation process for bioengineered foods. 

Canola seed is not commonly consumed without processing and its fractions have different uses for 

humans and livestock. Canola seed is processed into refined oil which is then used as a human food and 

livestock feed. The by-product meal fraction from processing is used as livestock feed. Refined oil is the 

only canola product that is consumed by humans. The oil is free from any protein and considered nonallergenic (Gylling, 2006; Hefle and Taylor, 1999; Moneret-Vautrin and Kanny, 2004), and therefore would 

not be expected to contain the GAT4621 protein. 

Human or livestock consumption of the GAT4621 protein in oil derived from 73496 canola will be 

negligible. Livestock will consume the GAT4621 protein in meal derived from 73496 canola seed after oil 

extraction. Any human exposure to GAT4621 protein would be limited to potential occupational 

exposure to canola seed meal and flour in feed processing facilities, grain mills or on farm. Due to the 

very low exposure and low risk, the GAT4621 protein in 73496 canola is safe. 

Although there will be consumption of the GAT4621 protein by livestock and also human occupational 

exposure from meal derived from 73496 canola, the GAT4621 protein is unlikely to cause an allergic 

reaction in humans or be a toxin in humans or animals, and therefore is safe for human exposure and 

animal consumption. 

1) The donor organism, B. licheniformis, is a common soil bacterium widely distributed in the 

environment. Due to its ubiquitous presence as spores in soil and dust, B. licheniformis is widely 

known as a contaminant of food but is not associated with any adverse effects. 

2) B. licheniformis has a history of safe use for the production of food enzymes in the United States, 

Canada, and Europe (e.g., alpha-amylase, cyclodextrin glycosyltransferase, hemicellulase, 

proteases, pullulanase), biocontrol agents (EU Commission, 2000; FDA, 2001) and as a probiotic 

(Alexopoulos et al., 2004a; Alexopoulos et al., 2004b; Kritas et al., 2006). B. licheniformis was 

determined by EPA to present low risk of adverse effects to human health and the environment and 

was subsequently granted an exemption for use in certain industrial fermentation processes (Federal 

Register, 1997). 

3) The amino acid sequence of the GAT4621 protein was compared to a database of allergens from the 

Food Allergy Research and Resource Program (FARRP), University of Nebraska, Allergen Database 

(FARRP, 2010; Version 10.0, January 2010; http://www.allergenonline.org/), which contains the amino 

acid sequences of 1471 known and putative allergenic proteins. Potential identities between the 

GAT4621 protein and proteins in the allergen database were evaluated using the FASTA34 sequence 

alignment program (Pearson and Lipman, 1988) set to the default parameters (ktup = 2, scoring matrix 

= BLOSUM50, gap creation penalty = -10, gap extension penalty = -2, E score cutoff = 10). The top 20 

high scoring alignments were reviewed for sequence identities greater than or equal to 35% over 80 or 

greater amino acid residues. None of the alignments met or exceeded the 35% over 80 or greater 

amino acid threshold. The GAT4621 amino acid sequence was also evaluated for any eight or greater 

contiguous identical amino acid matches to the same database of allergens noted above. There were 

no eight or greater contiguous identical amino acid matches observed with the GAT4621 amino acid 

sequence. These data indicate the lack of both amino acid identity and immunologically relevant 

similarities between the GAT4621 protein and known protein allergens. 

4) Additional experimental assays were conducted to assess the allergenic potential of the GAT4621 

protein. The GAT4621 protein was assayed for digestibility in simulated gastric fluid and in simulated 

intestinal fluids and also assayed for the presence of glycosylation. The GAT4621 protein is rapidly 

hydrolyzed in both simulated gastric and intestinal fluids (less than 30 seconds in simulated gastric 
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73496 Canola 

fluid containing pepsin at pH 1.2 as demonstrated by SDS-PAGE analysis, and less than 5 minutes in 

simulated intestinal fluid containing pancreatin at pH 7.5 as demonstrated by SDS-PAGE analysis). 

In addition, the GAT4621 protein in 73496 canola is not glycosylated (Appendix 4). Taken together 

with the similarity search results, these data indicate that the GAT4621 protein is unlikely to be 

allergenic to animals or humans. 

5) Bioinformatic analyses demonstrated that the GAT4621 protein retains the characteristics found in 

other N-acetyltransferases that are ubiquitous in plants and microorganisms (Neuwald and Landsman, 

1997). GAT4621 contains the definitive motif for the GNAT family of N-acetyltransferases (MarchlerBauer et al., 2005). This superfamily of enzymes is present in all organisms, including plants, 

mammals, fungi, algae, and bacteria (Dyda et al., 2000). 

6) An updated sequence similarity search of the GAT4621 protein sequence against the National Center 

for Biotechnology Information (NCBI) Entrez Protein dataset was conducted using the BLASTP 2.2.13 

algorithm against Release 176.0 (02/15/10) of the NCBI Entrez Protein dataset. A cutoff E score of 

1.0 was used to generate biologically meaningful similarity between the GAT4621 protein and 

proteins in the NCBI database, which allowed for identification of proteins with limited similarity in the 

search. Low complexity filtering was turned off and the maximum number of alignments returned was 

set to 2000. The GAT4621 similarity search identified 577 proteins that were within these criteria. 

None of the similar proteins returned by the search were identified as toxins, demonstrating that the 

GAT4621 protein is unlikely to share relevant sequence similarities with known protein toxins and is 

therefore unlikely to be a toxin itself. 

7) There was no evidence of acute toxicity in mice at a target dose of 2000 mg purified protein 

preparation per kg of body weight (equivalent to approximately 1640 mg of the full-length GAT4621 

protein per kg of body weight). Based on expression levels of GAT4621 protein in 73496 canola 

toasted meal, the highest calculated exposure to GAT4621 protein was for poultry livestock based on 

a hypothetical total replacement scenario. The margin of exposure for poultry to GAT4621 is greater 

than 50,000-fold the no observed adverse effect level of 1640 mg/ kg of body weight in mice. Based 

on these calculations, there is a wide margin of safety for the GAT4621 protein. 
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VII. Agronomic Performance and Ecological Observations

Agronomic and ecological evaluations were conducted to assess the comparability of 73496 canola to 

conventional canola. These evaluations form the basis to determine whether 73496 canola is 

comparable to conventional canola and is therefore no more likely to pose a plant pest risk than 

conventional canola. 

The agronomic evaluations were based on both laboratory experiments and replicated, multi-site field 

trials conducted by agronomists and scientists who are considered experts in the production and 

evaluation of canola. To evaluate the agronomic characteristics of 73496 canola, data were collected on 

representative characteristics that influence reproduction, crop survival, and potential weediness. In each 

of these assessments, 73496 canola was compared to a near-isoline control that was >95% genetically 

similar to 73496 canola but did not carry any recombinant DNA, and, in some experiments, was 

compared to non-transgenic commercial canola lines selected from current Pioneer conventional canola 

products. In each experiment, 73496 canola was comparable to the control or commercial comparators. 

The ecological evaluations included observed responses to insect and disease stressors during multiyear and multi-site field trials. These observations were made on 73496 canola and control canola and 

tracked the presence of insect and disease stressors in the field and the plant responses. In each case, 

73496 canola responded similarly to the control plants in these trials. 

Based on the analyses described below, 73496 canola is comparable to conventional canola and would 

not pose a greater plant pest risk or increased weed potential than conventional canola. 

VII-A. Germination Evaluations 

In order to test germination, seeds from the F1*5 generation (Section III-A, Figure 2) of 73496 canola were 

tested for germination under warm, cold, and diurnal conditions (Table 8). A near-isoline control was 

used for comparison. In addition, two commercial canola lines were evaluated to establish a reference 

range for germination. This range provided a context to determine the biological significance of any 

statistical differences observed. 

Each germination test contained eight replicates of 73496 canola, near-isoline control, and two 

commercial lines: one 60-seed replicate was counted and weighed for each canola line, with seven 

additional replicates weighed out based on the weight of the corresponding 60-seed replicate. The 

“Rules for Testing Seeds”, published by the Association of Official Seed Analysis, were used as 

guidelines for the germination methods and interpretation of results (AOSA, 2007). Each replicate was 

placed between sheets of moist germination toweling, rolled up, and placed in a growth chamber set to 

the appropriate test conditions as specified below. 

Table 8. Description of Seed Germination Conditions

Warm germination test  Continuous setting of 25°C and 90% relative humidity for 10 days 

 Germinated seed counted after 10 days 

Cold germination test 

 Continuous setting of 10°C and 90% relative humidity for 10 days, 

followed by three days at a continuous setting of 25°C and 90% relative 

humidity 

 Germinated seed counted after 13 days 

Diurnal germination test 

 Simulates daily weather conditions 

 Cyclical setting of 10°C and 90% relative humidity for 16 hours and then 

25°C and 90% relative humidity for 8 hours, repeated daily for 10 days 

 Germinated seed counted after 10 days 
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A statistically significant (P-value = 0.0460) difference in mean germination rate was observed between 

73496 canola and the control canola in the warm germination test, with a lower mean germination rate in 

73496 canola compared to the control canola (Table 9). However, for 73496 canola and the control 

canola, the range of germination rate values was within the reference range of commercial canola values. 

In the cold germination test, no statistically significant differences in mean germination rate were 

observed between 73496 canola and the control canola (Table 9). In the diurnal germination test, no 

statistically significant differences in mean germination rate were observed between 73496 canola and 

the control canola (Table 9). 

Table 9. Summary of Germination Results for 73496 Canola 

Statistic 73496 

Canola 

Control 

Canola 

Reference 

Rangea 

Warm n=525 n=469 

Meanb 96.4% 98.7% 

84% - 100% Rangec 93% - 100% 96% - 100% 

SEMd 0.00891 0.00535 

P-value 0.0460e

Cold n=518 n=466 

Mean 98.8% 98.4% 

88% - 100% Range 93% - 100% 97% - 100% 

SEM 0.00613 0.00745 

P-value 0.7127 

Diurnal n=514 n=453 

Mean 98.9% 99.8% 

92% - 100% Range 95% - 100% 98% - 100% 

SEM 0.00527 0.00224 

P-value 0.1635 

a

 Reference range denotes the lowest and highest germination rates across reference canola lines

b

 Mean percentage 

c

 Range denotes the lowest and highest germination rates 

d

 Standard error of the mean 

e

 Statistically significant difference; P-value <0.05 

Germination rates in 73496 canola under warm, cold and diurnal growing conditions were comparable to 

those of the control canola. A slightly lower germination rate was observed for 73496 canola seed under 

warm growing conditions that was statistically significant. However, this difference was small and the 

entire range of germination rate values for 73496 canola and the control canola under warm growing 

conditions were within the corresponding range of reference for commercial canola germination rates. 

The data provided here support the conclusion that 73496 canola is comparable to conventional canola 

with respect to germination. 

VII-B. Field Trial Evaluations 

73496 canola has been field tested in the United States since 2007 as authorized by USDA notification 

applications listed in Appendix 6. The list compiles a number of test sites in diverse regions of the U.S. 

including the major canola-growing areas. Agronomic data were collected to assess agronomic 

comparability as it relates to plant pest potential. In addition, extensive testing has also been conducted 

in Canada in major canola-growing areas, primarily because Pioneer’s canola breeding facilities are 

located in Canada and Canada is the primary market. A table of testing under authorized Canadian 

permits is also provided in Appendix 6. 
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Throughout the development process, 73496 canola was observed for unexpected differences in 

responses to abiotic stress (e.g, drought, excess moisture, temperature extremes, etc.). Monthly 

observations for response to naturally occurring abiotic stressors indicated that 73496 canola and nearisoline control were similar with respect to their response to abiotic stress. 

Agronomic data were collected from 73496 canola and control canola in two experiments (denoted A and 

B) that were conducted at 10 total field locations in 2008 and 2009 (Table 10; Figure 17). The trial 

locations were selected in order to provide data representative of the major canola growing regions in the 

U.S. and Canada, where commercial sales of 73496 canola are expected. Where sites were common 

between both experiments, the data from these sites were analyzed independently of each other. In 

addition, different characteristics were measured in each experiment because of the stage at which the 

data were collected. Experiment A was conducted earlier in the development of 73496 canola and was 

collected at Pioneer field sites, therefore the data were used as part of product advancement decisions. 

Experiment B was conducted at a later stage of development and at contractor sites, therefore the data 

were used to fully characterize the agronomic characteristics of 73496 canola. Agronomic practices used 

to prepare and maintain each field site were characteristic of each respective region. The genetic 

backgrounds selected for these trials are adapted for cultivation both in northern U.S. and in Canada. In 

addition, insect pests and diseases at the Canada locations are typical of those found in the U.S. While 

the majority of the sites were from Canadian field trial sites, these Canadian locations are considered 

representative of suitable canola growing areas within the U.S. Zone maps have been created by the 

U.S. EPA (EPA, 1996) and Canada Pest Management Regulatory Agency (PMRA, 1998) that divide 

North America into regions where climate conditions are similar (Figure 18, Panels A and B). These zone 

maps are used to determine where field trials should be placed when conducting studies for establishing 

tolerances (e.g. maximum pesticide residue levels) associated with domestic pesticide registrations. 

Thus, field trials conducted within the same zone are considered interchangeable. The zones for each of 

the field trials conducted are provided in Table 10 and show that the zone designations for trials 

conducted in Canada are representative of U.S. zones. 

A statistical analysis of agronomic data was conducted to test for differences in the mean values between 

the 73496 canola and the near-isoline control (Appendix 2 for statistical model). When numerous 

comparisons are being made, it is important to control the rate of false positive results. Since the 

introduction of the false discovery rate (FDR) approach in the mid-1990’s, it has been widely employed 

across a number of scientific disciplines, including genomics, ecology, medicine, plant breeding, 

epidemiology, dairy science and signal/image processing (e.g., Pawitan et al., 2005; Spelman and 

Bovenhuis, 1998). A false positive result occurs when two means are deemed significantly different 

when, in fact, they are not. If one uses a 5% type I error rate for each agronomic characteristic 

measured, then the number of false positives increases as the number of characteristics increase. In 

order to help manage the false positive rate, the FDR method of Benjamini and Hochberg was applied to 

account for making multiple comparisons (Benjamini and Hochberg, 1995; Westfall et al., 1999). 

P-values were adjusted accordingly. This resulted in the false positive rate being held to 5%. Both 

adjusted and unadjusted P-values are provided for the agronomic data.
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1

2

3, 9, 10

4 5

6

7

8

Canada

United States

 Figure 17. Map of Locations for Agronomic Data Collection for 73496 Canola 

 Table 10. Site Locations for 2008 and 2009 Field Trial Experiments 

Map Site 

Number Country Location Zone 

Experiment A: 2008 Field Trials 

1 

Canada 

Riviere Qui Barre, AB 14 

2 Nisku, AB 14 

3 Saskatoon, SK 7/14 

4 Crystal City, MB 5/14 

5 Rosebank, MB 5/14 

6 Morden, MB 5/14 

7 Georgetown, ON 5 

Experiment B: 2009 Field Trials 

8 U.S. Ephrata, WA 11 

9 Canada Saskatoon, SK 7/14 

10 Saskatoon, SK 7/14 
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A. Canada PMRA Zones 

B. U.S. EPA Zones 

Figure 18. Maps Indicating Zones with Similar Growing Conditions for Field Trials 

Panel A: Zones established by PMRA in Canada with similar climate conditions for residue field trials (PMRA, 

1998). Panel B: Zones established by the U.S. EPA with similar climate conditions for residue field trials (EPA, 

1996).
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73496 Canola 

Experiment A contains data from seven locations planted in Canada during the 2008 growing season 

(Table 10, Figure 17). The purpose of Experiment A was to evaluate the agronomic characteristics and 

yield of 73496 canola. Seed of the T3 generation of 73496 canola was used (Section III-A, Figure 2). 

The control plants were near-isoline control canola. 

The following characteristics were measured: early growth, plant height, days to flower, days to maturity, 

and yield. Descriptions of the characteristics and their measurement are found in Table 11. 

Each site included a randomized complete block design containing three blocks, with each block 

containing one plot of 73496 canola and one plot of the control canola. Procedures employed to control 

the introduction of experimental bias in this study were as follows: non-systematic selection of trial areas 

and plot areas within each site, randomization of the canola entries within each block, and uniform 

maintenance treatments across each plot area. 

Results of Experiment A are summarized in Table 12. 

For characteristics early growth, plant height, days to maturity, and yield, no statistically significant 

differences were observed between the mean values of 73496 canola and the control. One statistical 

difference was observed between 73496 canola and the control in days to flower (adjusted P-value < 

0.05). The mean flowering time was one day longer for 73496 canola; however, the range of individual 

values for days to flower in 73496 canola was within the range of the control data. Furthermore, this 

statistical difference was not replicated in the trial conducted in 2009. Because the time to flowering for 

73496 canola is still within the normal range of canola flowering, this statistical difference is not 

biologically meaningful to the determination of plant pest potential. 

The results from Experiment A indicate that 73496 canola is agronomically comparable to the near-isoline 

control. 

Table 11. Experiment A: 2008 Field Trial Agronomic Characteristics Measured 

General 

Characteristic 

Characteristic 

Measured 

Number 

of Sites Data Description Scale 

Vegetative 

Parameter 

Early Growth 3 of 7 

Overall plant health and 

amount of soil surface area 

covered by leaves at the 4-leaf 

to 6-leaf stages 

1-9 Scale where 

1=Unhealthy/weak looking 

plants with small leaf 

coverage; 

9=healthy/strong-looking 

plants with large leaf 

coverage 

Plant Height 7 of 7 

10 plants per plot at 

physiological maturity. Height 

was measured from the soil 

surface to the tip of the highest 

pod. 

Centimeters 

Reproductive 

Parameter 

Days to Flower 7 of 7 

Number of days from the 

planting date to the date when 

approximately 50% of plants 

had produced flowers. 

Days 

Days to Maturity 6 of 7 

Number of days from the 

planting date to the date when 

approximately 50% of plants 

had reached physiological 

maturity. 

Days 

Yield 7 of 7 

Weight (corrected for moisture 

content of seed) harvested 

from all plants in a given plot 

kg/ha 
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Table 12. Experiment A: Summary of Agronomic Characteristics of 73496 Canola Across Seven 

Locations 

Agronomic Characteristica

(unit of measurement) 

Control 

Canola 

73496 

Canola 

Early Growth 

(1-9 scale) 

n=9 

Meanb

 6.50 7.37 

Rangec 6 - 7 6 - 8 

Standard Error 0.445 0.445 

P-Valued 0.0395 

Adjusted P-Valuee 0.0790 

Days to Flower 

(days) 

n=21 

Mean 44.6 45.7 

Range 40 - 50 42 - 49 

Standard Error 1.09 1.09 

P-Value 0.000218 

Adjusted P-Value 0.00131f

Days to Maturity 

(days) 

n=18 

Mean 93.0 92.8 

Range 87 - 106 89 - 105 

Standard Error 2.35 2.35 

P-Value 0.468 

Adjusted P-Value 0.562 

Plant Height 

(centimeters) 

n=21 

Mean 89.5 91.6 

Range 65 - 115 60 - 118 

Standard Error 5.10 5.10 

P-Value 0.355 

Adjusted P-Value 0.532 

Yield 

(kg/ha) 

n=21 

Mean 2310 2140 

Range 1020-3500 1140-2910 

Standard Error 199 199 

P-Value 0.0311 

Adjusted P-Value 0.0790 a Refer to Table 11 for descriptions of each agronomic characteristic measured. b Least squares mean c Range denotes the lowest and highest individual value across sites. d Non-adjusted P-value e False Discovery Rate (FDR) adjusted P-value f Statistically significant difference, adjusted P-value <0.05 
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73496 Canola 

Experiment B contains data from three locations planted in the U.S. and Canada during the 2009 

growing season (Table 10, Figure 17). The purpose of Experiment B was to evaluate the agronomic 

characteristics and yield of 73496 canola. Seed from the T5 generation was used (Section III-A, Figure 

2). The control plants were near-isoline control canola 

The following agronomic characteristics were measured in each block: early population, seedling vigor, 

days for flowering, plant height, disease incidence, insect damage, lodging, days to maturity, shattering, 

final population, and yield data. Descriptions of the characteristics and their measurement are found in 

Table 13. 

Each site included a randomized complete block design containing four blocks, with each block 

containing 73496 canola and control canola. 

Results of Experiment B are summarized in Table 14. 

No statistically significant differences were observed between 73496 canola and control canola mean 

values for early population, final population, seedling vigor, days to flowering, flowering duration, disease 

incidence, insect damage, lodging, days to maturity, shattering, yield, or plant height. 

The results from Experiment B indicate that 73496 canola is agronomically comparable to the near-isoline 

control. 

Table 13. Experiment B: 2009 Field Trial Agronomic Characteristics Measured 

General 

Characteristic 

Characteristic 

Measured 

Evaluation 

timinga Data Description Scale 

Germination / 

Emergence 

Early Population BBCH 11-13 

Number of plants emerged 

per plot (two 1 square meter 

quadrants) during true leaf 

(rosette) development 

Actual count per 

plot 

Seedling Vigor BBCH 11-13 

Visual estimate of average 

vigor of emerged plants per 

plot during true leaf (rosette) 

development 

From 1 to 9, where 

1=short plants with 

small thin leaves, 

and 

9=tall plants with 

large robust leaves 

Vegetative 

Parameters 

Plant Height BBCH 79 

Height from the soil surface 

to the tip of the highest pod 

when nearly all pods on the 

plants are at full size 

Height in cm 

Lodging BBCH 87-89 

Visual estimate of lodging 

severity when >70% of pods 

are fully ripe 

From 1 to 9, where 

1=plants lying flat, 

and 

9=plants standing 

straight 

Shattering BBCH 87-89 

Visual estimate of seed 

shattering when >70% of 

pods are fully ripe 

From 1 to 9, where 

1=high seed 

shattering, and 

9=no seed 

shattering 

Final Population Harvest Total number of plants per 

plot 

Actual count per 

plot 

a

 Based on Lancashire et al. (1991) for canola growth stages. 
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Table 13. Experiment B: 2009 Field Trial Agronomic Characteristics Measured (continued)

General 

Characteristic 

Characteristic 

Measured 

Evaluation 

timinga Data Description Scale 

Reproductive 

Parameters 

Days to 

flowering BBCH 57-62 From the time of planting to 

the first open flowers. Days 

Flowering 

duration BBCH 69 

Number of days from the 

date when the first flowers 

were open to the date 

when flowering ended on 

approximately 50% of the 

plants. 

Days 

Days to Maturity BBCH 86-87 

Number of days from 

planting to the date when 

between 60-70% of pods 

are ripe. 

Days 

Yield At harvest 

Weight (corrected for 

moisture content of seed) 

harvested from all plants in 

a given plot 

kg/ha 

Ecological 

Interactions 

Disease 

Incidence BBCH 71-79 

Visual estimate of foliar 

disease incidence during 

plant pod elongation. 

Ranging from 1-9 

where 

1 = poor disease 

resistance or high 

infection; 9 = best 

disease resistance 

or low infection. 

Insect Damage BBCH 71-79 

Visual estimate of insect 

damage during plant pod 

elongation. 

Ranging from 1-9 

where 

1 = poor insect 

resistance or high 

damage; 

9 = best insect 

resistance or low 

damage. a

 Based on Lancashire et al. (1991) for canola growth stages. 
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73496 Canola 

Table 14. Experiment B: Summary of Agronomic Characteristics of 73496 Canola Across Three 

Locations 

Agronomic Characteristica

(unit of measurement) 

Control 

Canola 

73496 

Canola 

Early Population 

(number of plants) 

73496 Canola n=11 

Control Canola n=12

Meanb 108 116 

Rangec 50 - 184 41 - 207 

CId, e (24) - 239 (16) - 247 

P-Valuef 0.496 

Adjusted P-Valueg 0.661 

Final Population 

(number of plants) 

73496 Canola n=11 

Control Canola n=12

Mean 87 82 

Range 29 - 158 23 - 164 

CI (9) - 183 (14) - 177 

P-Value 0.691 

Adjusted P-Value 0.691 

Seedling Vigor 

(1-9 scale) 

73496 Canola n=11 

Control Canola n=12

Mean 5 5 

Range 3 - 8 3 - 8 

CI 0 - 11 0 - 11 

P-Value 0.422 

Adjusted P-Value 0.632 

Days to Flowering 

(number of days) 

73496 Canola n=11 

Control Canola n=12

Mean 47 47 

Range 35 - 60 35 - 59 

CI 17 - 76 18 - 77 

P-Value 0.327 

Adjusted P-Value 0.561 

Flowering Duration 

(number of days) 

73496 Canola n=11 

Control Canola n=12

Mean 27 28 

Range 22 - 37 22 - 38 

CI 8 - 47 8 - 48 

P-Value 0.116 

Adjusted P-Value 0.279 

Disease Incidence 

(1-9 scale) 

73496 Canola n=11 

Control Canola n=12

Mean 7 7 

Range 6 - 9 6 - 8 

CI 5 - 10 5 - 9 

P-Value 0.0851 

Adjusted P-Value 0.279 

Insect Damage 

(1-9 scale) 

73496 Canola n=11 

Control Canola n=12

Mean 6 7 

Range 3 - 9 3 - 9 

CI (1) - 14 (1) - 14 

P-Value 0.165 

Adjusted P-Value 0.329 

Lodging 

(1-9 scale) 

73496 Canola n=11 

Control Canola n=12

Mean 8 8 

Range 7 - 9 6 - 9 

CI 6 - 10 6 - 10 

P-Value 0.624 

Adjusted P-Value 0.691 a Refer to Table 13 for descriptions of each agronomic characteristic measured. b Least squares mean c Range denotes the lowest and highest individual value across sites for 73496 canola and near-isoline control. d

 Confidence interval 

e

 Confidence intervals within parentheses indicate a negative value.

f Non-adjusted P-value 

g False Discovery Rate (FDR) adjusted P-value 
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Table 14. Experiment B: Summary of Agronomic Characteristics of 73496 Canola Across Three 

Locations (continued) 

Agronomic Characteristica

(unit of measurement) 

Control 

Canola 

73496 

Canola 

Days to Maturity 

(numbers of days) 

73496 Canola n=11 

Control Canola n=12

Meanb

 113 114 

Rangec 103 - 132 104 - 133 

CId,e 73 - 152 74 - 153 

P-Valuef 0.102 

Adjusted P-Valueg 0.279 

Shattering 

(1-9 scale) 

73496 Canola n=11 

Control Canola n=12

Mean 8 7 

Range 5 - 9 4 – 9 

CI 5 - 10 5 - 10 

P-Value 0.646 

Adjusted P-Value 0.691 

Yieldh

(kg/ha at 8% moisture) 

73496 Canola n=8 

Control Canola n=8

Mean 3500 2900 

Range 2600 - 4680 1550 - 4280 

CI 1620 - 5380 1030 - 4780 

P-Value 0.0395 

Adjusted P-Value 0.279 

Plant Height 

(centimeters) 

73496 Canola n=110 

Control Canola n=120

Mean 118 113 

Range 100 - 143 94 - 134 

CI 81 - 155 76 - 150 

P-Value 0.0563 

Adjusted P-Value 0.279 a Refer to Table 13 for descriptions of each agronomic characteristic measured. b Least squares mean c Range denotes the lowest and highest individual value across sites for 73496 canola and near-isoline control. d

 Confidence interval 

e

 Confidence intervals within parentheses indicate a negative value.

f Non-adjusted P-value 

g False Discovery Rate (FDR) adjusted P-value h Yield data from one of the three sites was not included due to an early killing frost that affected seed harvest at 

maturity. 

VII-C. Ecological Observations 

Ecological observations (i.e., plant interactions with pest insects and diseases) were recorded for all 

USDA-APHIS permitted field trials of 73496 canola during the 2007, 2008, 2009, and 2010 growing 

seasons. In addition, trials were conducted in Canada in 2007, 2008, and 2009 and the ecological 

observations were recorded. Plant breeders and field staff familiar with plant pathology and entomology 

observed 73496 canola and control lines at least every four weeks for pest insect and disease pressure 

and recorded the severity of any stressor seen. Any unexpected differences in response between 73496 

canola and various control lines (near-isolines and/or conventional canola lines) were recorded. 

The following scale was used when recording observations: 

 mild – very little disease or insect injury (<10%) visible 

 moderate – noticeable plant tissue damage (10% to 30%) 

 severe – significant plant tissue damage (>30%) 

A summary of the insect and disease ecological observations for both U.S. and Canada field trials is 

provided in Appendix 5. In every case, the severity of insect or disease stress on 73496 canola was not 

qualitatively different from various control lines growing at the same location. These results support the 

conclusion that the ecological interactions for 73496 canola were comparable to control canola lines with 

similar genetics or to conventional canola lines. 
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73496 Canola 

VII-D. Conclusions on Agronomic Performance and Ecological Observations 

73496 canola was observed in laboratory experiments and at 10 field locations in the U.S. and Canada to 

measure agronomic parameters. Data generated from these studies represent observations that are 

typically recorded by plant breeders and agronomists to evaluate the characteristics of canola over a 

broad range of environmental conditions that represent where 73496 canola may be grown. The 

agronomic characteristics measured are representative of reproduction, survival, and potential 

weediness. The measured characteristics provide data useful in establishing a basis to assess 

agronomic comparability and familiarity of 73496 canola compared to conventional canola in the context 

of ecological risk assessment. 

The agronomic data showed no statistical differences between 73496 canola and control canola 

(untransformed near-isoline controls and/or commercial canola lines) except for days to flower in 

Experiment A. The range of individual values for days to flower in 73496 canola was within the range of 

the control data and the statistical difference was not replicated in Experiment B. These data support the 

conclusion that 73496 canola is comparable in agronomic characteristics to conventional canola. 

Ecological observations of U.S. and Canadian field trials for responses of 73496 canola to naturally 

occurring insect and disease stressors showed no unexpected differences from control canola. The 

assessment of the ecological data detected no biologically significant differences between 73496 canola 

and control canola lines indicative of a selective advantage that would result in increased weed potential 

or plant pest risk. 

Based on these analyses, 73496 canola is comparable to conventional canola and would not pose a 

greater plant pest risk or increased weed potential than conventional canola. 
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73496 Canola 

VIII. Compositional Assessment

Compositional comparisons between transgenic crops and conventional varieties are a key part of a 

nutritional and safety assessment and provide assurance of the food safety of transgenic crops. 

Compositional assessments are performed in accordance with the principles outlined in the OECD 

consensus document on compositional considerations for canola (OECD, 2001). This document 

emphasizes quantitative measurements of essential nutrients, and known anti-nutrients and toxicants. 

These analyses will effectively highlight any compositional changes that may indicate potential safety and 

anti-nutritional concerns. 

For 73496 canola, the following analytes were measured for the comparative assessment: proximates, 

fiber, fatty acids, amino acids, vitamins, minerals, glucosinolates, secondary metabolites, phytosterols, 

and anti-nutrients. Levels of the analytes were measured in the seed of the F1*4 generation (Section III-A; 

Table 1, Figure 2) of 73496 canola and were compared to corresponding levels in the near-isoline control 

and were also compared to statistical tolerance intervals generated from non-modified conventional 

commercial varieties. These comparisons formed the basis for determining compositional comparability 

of 73496 canola to conventional canola. Canola seed was chosen as the test material for the 

compositional analysis of 73496 canola because oil fractions are derived from seed. Compositional 

evaluation of seed would be representative of these derived materials. 

Seed samples were collected from five separate sites in Canada and the United States during the 2009 

growing season (three sites in Manitoba, Canada: Elm Creek, Minto, and Portage la Prairie; two sites in 

the United States: Velva, ND and Ephrata, WA). Each site utilized a randomized complete block design 

with four blocks and each block containing 73496 canola and the control canola. Each plot of 73496 

canola was treated with glyphosate at a rate of 0.367-0.414 pounds acid equivalent per acre (411-464 

grams acid equivalent per hectare) consistent with typical agronomic practices, and each plot of the 

control canola was left untreated. Seed samples collected from 73496 canola and the control canola 

were analyzed for key nutritional components in accordance with the OECD consensus document on 

compositional considerations for new varieties of canola (OECD, 2001), which included analysis of 

proximates, fiber, fatty acids, amino acids, vitamins, minerals, glucosinolates, secondary metabolites, 

phytosterols, and anti-nutrients. 

In a separate experiment for the statistical tolerance intervals, seed was also collected from five 

conventional (i.e., non-modified) commercial canola lines grown at five field locations in canola-growing 

areas of U.S. and Canada in 2008 and 2009. Planting, harvesting, processing, and compositional 

analysis procedures for the reference hybrid trials were similar to those employed for the trials containing 

near-isoline control and 73496 canola. Compositional analyses of the reference lines were used to 

determine the statistical tolerance intervals, which established the normal variation for the measured 

analytes in canola. 

Statistical analysis of nutrient composition data was conducted to test for differences in the analyte mean 

values between 73496 canola and the near-isoline control. For details of the statistical methodology, 

refer to Appendix 2. When numerous analytes are being evaluated on the same samples, controlling 

false positive outcomes is important. A false positive outcome occurs when an analyte mean of the 

transgenic line is deemed significantly different from the analyte mean of the control line, when in fact the 

two means are not different. If one uses a 5% type I error rate for each analyte, then the number of false 

positives increases as the number of analytes increase. In order to help manage the false positive rate, 

the false discovery rate (FDR) method of Benjamini and Hochberg was applied to account for making 

multiple comparisons (Benjamini and Hochberg, 1995; Westfall et al., 1999). P-values were adjusted 

accordingly, resulting in the false positive rate being held to 5%. Both adjusted and non-adjusted 

P-values are provided in this submission. In the discussion of these data, a significant difference 

between the mean of 73496 canola and that of the near isoline was established with an FDR-adjusted Pvalue <0.05. 

Using the data obtained from the commercial lines, a statistical tolerance interval was calculated to 

contain, with 95% confidence, 99% of the values contained in the population of commercial canola. Any 
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73496 Canola 

negative limits were set to zero. This statistical tolerance interval provided further context for 

interpretation of the composition results for 73496 canola. 

VIII-A. Proximate and Fiber Analysis 

Analysis of the major constituents of canola seed, or proximates, was used to determine the nutritional 

properties of seed from 73496 and control canola. No statistically significant differences were observed 

between 73496 canola and the control canola mean values for any of the proximate and fiber analytes 

(Table 15). 

In conclusion, proximate and fiber analysis of canola seed demonstrates that 73496 canola is comparable 

to conventional canola.
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73496 Canola 

Table 15. Summary Analysis of Proximate and Fiber Results in Canola Seed 

Analyte 

Control 

Canola 

n=20 

73496 

Canola 

n=20 

Tolerance 

Intervala

% Dry Weight

Crude Protein 

Meanb 26.6 25.9 

11.3 - 54.4 

Rangec 23.4 - 31.9 22.4 - 31.6 

CId 23.4 - 30.3 22.8 - 29.5 

P-Valuee 0.0275 

Adjusted P-Valuef 0.0974 

Crude Fat 

Mean 43.6 43.9 

30.6 - 65.8 

Range 37.4 - 51.5 37.0 - 49.6 

CI 40.1 - 47.3 40.3 - 47.7 

P-Value 0.608 

Adjusted P-Value 0.737 

ADFg

Mean 32.4 32.3 

14.5 - 63.4 

Range 27.1 - 38.6 26.6 - 39.5 

CI 28.8 - 36.5 28.7 - 36.4 

P-Value 0.929 

Adjusted P-Value 0.957 

Crude Fiber 

Mean 28.4 28.5 

12.9 - 55.5 

Range 24.3 - 35.1 22.8 - 35.5 

CI 25.1 - 32.1 25.2 - 32.2 

P-Value 0.889 

Adjusted P-Value 0.935 

NDFh

Mean 33.4 33.3 

15.5 - 67.7 

Range 28.8 - 40.3 28.0 - 38.9 

CI 30.4 - 36.8 30.2 - 36.7 

P-Value 0.844 

Adjusted P-Value 0.911 

Ash 

Mean 4.08 3.92 

2.01 - 7.63 

Range 2.98 - 5.10 3.00 - 5.06 

CI 3.29 - 5.07 3.16 - 4.87 

P-Value 0.0549 

Adjusted P-Value 0.141 

Carbohydrates 

Mean 25.3 26.0 

19.6 - 33.5 

Range 16.9 - 29.1 23.7 - 28.8 

CI 24.0 - 26.7 24.6 - 27.4 

P-Value 0.320 

Adjusted P-Value 0.468 a The statistical tolerance interval was calculated from commercial canola varieties and calculated to contain with 95 

percent confidence, 99 percent of the population of canola, negative limits set to zero b Least squares mean were generated from a mixed model analysis c Range denotes the lowest and highest individual value across sites d

 Confidence Interval 

e Non-adjusted P-value f

 False Discovery Rate (FDR) adjusted P-value g

 Acid Detergent Fiber 

h

 Neutral Detergent Fiber 
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73496 Canola 

VIII-B. Fatty Acid Analysis 

Five fatty acids account for more than 96 percent of the total fatty acids in canola seed, with the most 

abundant being oleic (C18:1 Δ9; ~ 60%) and linoleic (C18:2 Δ9,12; ~ 20%) acids. Less abundant, but 

occurring at measurable concentrations are palmitic (C16:0), stearic (C18:0) and α-linolenic (C18:3 

Δ9,12,15) acids. The desaturation of oleic acid to form linoleic acid, and its subsequent desaturation to 

form α-linolenic acid, occurs only in plants, hence both linoleic and α-linolenic acids are essential fatty 

acids for mammals. For this reason, it was important to analyze any unintended changes in the 

concentrations of linoleic and α-linolenic acids, and their key precursors palmitic, stearic, and oleic acids, 

in seed from 73496 canola. 

Other polyunsaturated fatty acids, such as γ-linolenic (C18:3 Δ6,9,12), eicosatrienoic (C20:3 Δ8,11,14) 

and arachidonic (C20:4 Δ5,8,11,14) acids can all be synthesized by mammals from dietary sources of 

α-linolenic and linoleic acid. Hence, small changes in the concentrations of these trace fatty acids in 

73496 canola seed would have little or no biological significance to either humans or animals consuming 

derived products. Similarly, the synthesis of palmitoleic (C16:1 Δ9) and saturated fatty acids with chain 

lengths greater than 18 (e.g., C20:0, C22:0, C24:0), can be accomplished in mammals through de novo

fatty acid synthesis without dietary requirements for palmitic and stearic acids, respectively. 

The fatty acids reported in Table 16 were identified by OECD as those that should be measured for 

compositional comparisons in canola (OECD, 2001). Based on consultation with the Codex Alimentarius 

Commission definition of canola oil (CODEX, 2005), six additional fatty acids were reported in this 

analysis. Non-detectable fatty acids are defined as <0.05% of total fatty acids (CODEX, 2005). 

Heptadecanoic acid (C17:0), γ-linolenic acid (C18:3), and erucic acid (C22:1) were below this threshold 

and were not included in Table 16. As expected per the canola definition, erucic acid (C22:1) was not 

detected in the analysis. 

Small but statistically significant differences were observed using the adjusted P-value in concentrations 

of oleic and linoleic acid between 73496 and control canola (Table 16). However, these fatty acid 

concentrations were within the respective tolerance intervals derived for commercial canola varieties. 

Therefore, the small, statistically significant differences observed for certain fatty acids are unlikely to be 

biologically meaningful. 

In conclusion, fatty acid analysis of canola seed demonstrates that 73496 canola is comparable to 

conventional canola. 
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73496 Canola 

Table 16. Summary Analysis of Fatty Acid Composition in Canola Seed 

Analyte 

Control 

Canola 

n=20 

73496 

Canola 

n=20 

Tolerance

Intervala

% Total Fatty Acids

Lauric Acid (C12:0) 

Meanb 0.0816 0.0811 

0 - 0.216 

Rangec 0.0568 - 0.105 0.0563 - 0.113 

CId 0.0639 - 0.104 0.0634 - 0.104 

P-Valuee 0.863 

Adjusted P-Valuef 0.919 

Myristic Acid (C14:0) 

Mean 0.053 0.0546 

0.0239 - 

0.112 

Range 0.0470 - 0.0749 0.0484 - 0.0670 

CI 0.0466 - 0.0603 0.0480 - 0.0621 

P-Value 0.212 

Adjusted P-Value 0.355 

Palmitic Acid (C16:0) 

Mean 4.08 4.19 

3.10 - 5.60 

Range 3.72 - 4.40 3.85 - 4.45 

CI 3.87 - 4.31 3.97 - 4.42 

P-Value 0.0111 

Adjusted P-Value 0.0605 

Palmitoleic Acid (C16:1) 

Mean 0.262 0.272 

0.154 - 

0.378 

Range 0.225 - 0.306 0.224 - 0.317 

CI 0.232 - 0.295 0.241 - 0.307 

P-Value 0.0173 

Adjusted P-Value 0.079 

Heptadecenoic Acid (C17:1) 

Mean 0.108 0.105 

0 - 0.264 

Range 0.0675 - 0.140 0.0666 - 0.137 

CI 0.0802 - 0.146 0.0776 - 0.141 

P-Value 0.058 

Adjusted P-Value 0.144 

Stearic Acid (C18:0) 

Mean 1.71 1.61 

1.17 - 3.47 

Range 1.47 - 1.89 1.44 - 1.93 

CI 1.57 - 1.86 1.48 - 1.76 

P-Value 0.00577 

Adjusted P-Value 0.0507 

Oleic Acid (C18:1) 

Mean 61.6 60.1 

53.1 - 74.4 

Range 58.6 - 64.1 56.8 - 63.3 

CI 58.9 - 64.4 57.5 - 62.9 

P-Value <0.0001 

Adjusted P-Value 0.00216g

Linoleic Acid (C18:2) 

Mean 20.1 21.3 

13.1 - 27.1 

Range 18.5 - 22.5 19.5 - 23.4 

CI 18.5 - 21.8 19.5 - 23.1 

P-Value 0.000107 

Adjusted P-Value 0.00220g

(9,15) Isomer of Linoleic Acid 

(C18:2) 

Mean 0.0986 0.0924 

0 - 0.524h

Range 0.0626 - 1.45 0.0611 - 0.496 

CI 0.0697 - 0.139 0.0653 - 0.131 

P-Value 0.764 

Adjusted P-Value 0.868 
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73496 Canola 

Table 16. Summary Analysis of Fatty Acid Composition in Canola Seed (continued) 

Analyte 

Control 

Canola 

n=20 

73496 

Canola 

n=20 

Tolerance 

Interval 

% Total Fatty Acids

Linolenic Acid (C18:3) 

Mean 8.92 9.29 

4.23 - 16.8 

Range 7.46 - 10.0 8.08 - 10.5 

CI 8.04 - 9.89 8.38 - 10.3 

P-Value 0.00884 

Adjusted P-Value 0.0525 

Arachidic Acid (C20:0) 

Mean 0.615 0.586 

0.496 - 1.01 

Range 0.585 - 0.645 0.552 - 0.640 

CI 0.598 - 0.632 0.570 - 0.602 

P-Value 0.00618 

Adjusted P-Value 0.0507 

Eicosenoic Acid (C20:1) 

Mean 1.38 1.36 

0.999 - 1.88 

Range 1.19 - 1.54 1.24 - 1.45 

CI 1.27 - 1.51 1.25 - 1.48 

P-Value 0.128 

Adjusted P-Value 0.269 

Eicosadienoic Acid 

(C20:2) 

Mean 0.069 0.0713 

0.0308 - 

0.107 

Range 0.0565 - 

0.0805 

0.0563 - 

0.0831 

CI 0.0583 - 

0.0817 

0.0602 - 

0.0844 

P-Value 0.0256 

Adjusted P-Value 0.0974 

Behenic Acid (C22:0) 

Mean 0.328 0.319 

0.241 - 

0.465 

Range 0.290 - 0.365 0.286 - 0.359 

CI 0.300 - 0.358 0.292 - 0.348 

P-Value 0.0182 

Adjusted P-Value 0.079 

Lignoceric Acid (C24:0) 

Mean 0.178 0.17 

0.0601 - 

0.657 

Range 0.160 - 0.208 0.151 - 0.206 

CI 0.160 - 0.198 0.153 - 0.189 

P-Value 0.0183 

Adjusted P-Value 0.079 

Nervonic Acid (C24:1) 

Mean 0.191 0.194 

0.0295 - 

0.542 

Range 0.121 - 0.257 0.120 - 0.254 

CI 0.141 - 0.259 0.143 - 0.263 

P-Value 0.303 

Adjusted P-Value 0.458 a The statistical tolerance interval was calculated from commercial canola varieties and calculated to contain with 95 

percent confidence, 99 percent of the population of canola, negative limits set to zero b Least squares mean were generated from a mixed model analysis c Range denotes the lowest and highest individual value across sites d Confidence Interval 

e Non-adjusted P-value f False Discovery Rate (FDR) adjusted P-value g

 Statistically significant difference; FDR Adjusted P-Value <0.05 h Tolerance interval could not be calculated (NC) due to all sample values below the assay LLOQ. 
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73496 Canola 

VIII-C. Amino Acid Analysis 

Canola seed is generally a good source of essential and non-essential amino acids for most domestic 

animal species. Total levels of 18 amino acids and 26 free amino acids were measured in 73496 and 

control seed. 

Results are shown in Table 17 for the 18 total amino acids. No statistically significant differences 

between the 73496 canola and control were noted for any amino acids measured (adjusted P-value). 

The levels of 26 free amino acids were also measured (Pioneer data not shown). For each analyte, the 

measured values for 73496 canola were within statistical tolerance intervals defined by commercial 

conventional varieties even when a statistically significant difference was observed between 73496 

canola and control. 

In conclusion, total and free amino acid analysis of canola seed demonstrates that 73496 canola is 

comparable to conventional canola. 
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73496 Canola 

Table 17. Summary Analysis of Amino Acid Composition in Canola Seed 

Analyte 

Control 

Canola 

n=20 

73496 

Canola 

n=20 

Tolerance 

Intervala

% Dry Weight

Alanine 

Meanb 1.14 1.12 

0.588 - 1.90 

Rangec 1.01 - 1.37 0.970 - 1.32 

CId 1.02 - 1.28 0.996 - 1.25 

P-Valuee 0.0446 

Adjusted P-Valuef 0.133 

Arginine 

Mean 1.62 1.57 

0.741 - 3.07 

Range 1.30 - 1.98 1.32 - 1.92 

CI 1.42 - 1.85 1.37 - 1.79 

P-Value 0.0502 

Adjusted P-Value 0.133 

Aspartic Acid 

Mean 2.00 2.06 

0.980 - 3.52 

Range 1.26 - 2.54 1.77 - 2.50 

CI 1.71 - 2.33 1.77 - 2.40 

P-Value 0.152 

Adjusted P-Value 0.312 

Cystine 

Mean 0.606 0.618 

0.311 - 1.24 

Range 0.505 - 0.751 0.523 - 0.722 

CI 0.516 - 0.712 0.526 - 0.726 

P-Value 0.592 

Adjusted P-Value 0.737 

Glycine 

Mean 1.38 1.35 

0.688 - 2.52 

Range 1.09 - 1.61 1.17 - 1.54 

CI 1.25 - 1.52 1.22 - 1.49 

P-Value 0.162 

Adjusted P-Value 0.317 

Glutamic Acid 

Mean 5.05 5.01 

1.99 - 11.9 

Range 2.48 - 6.68 4.30 - 6.40 

CI 4.23 - 6.02 4.20 - 5.97 

P-Value 0.783 

Adjusted P-Value 0.868 

Histidine 

Mean 0.800 0.801 

0.342 - 1.72 

Range 0.644 - 0.966 0.667 - 0.939 

CI 0.712 - 0.899 0.713 - 0.900 

P-Value 0.945 

Adjusted P-Value 0.957 

Isoleucine 

Mean 1.08 1.06 

0.533 - 1.95 

Range 0.869 - 1.30 0.922 - 1.25 

CI 0.958 - 1.22 0.935 - 1.19 

P-Value 0.113 

Adjusted P-Value 0.250 

Leucine 

Mean 1.88 1.83 

0.910 - 3.42 

Range 1.52 - 2.26 1.59 - 2.19 

CI 1.66 - 2.12 1.62 - 2.07 

P-Value 0.0498 

Adjusted P-Value 0.133 
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73496 Canola 

Table 17. Summary Analysis of Amino Acid Composition in Canola Seed (continued) 

Analyte 

Control 

Canola 

n=20 

73496 

Canola 

n=20 

Tolerance 

Interval 

% Dry Weight 

Lysine 

Mean 1.65 1.64 

0.729 - 3.16 

Range 1.25 - 2.02 1.48 - 1.97 

CI 1.45 - 1.87 1.45 - 1.86 

P-Value 0.669 

Adjusted P-Value 0.793 

Methionine 

Mean 0.464 0.472 

0.258 - 0.857 

Range 0.383 - 0.546 0.402 - 0.546 

CI 0.408 - 0.529 0.415 - 0.538 

P-Value 0.596 

Adjusted P-Value 0.737 

Phenylalanine 

Mean 1.12 1.10 

0.545 - 2.12 

Range 0.901 - 1.35 0.927 - 1.28 

CI 1.01 - 1.26 0.985 - 1.23 

P-Value 0.167 

Adjusted P-Value 0.319 

Proline 

Mean 1.63 1.59 

0.717 - 3.36 

Range 1.46 - 1.98 1.38 - 1.95 

CI 1.43 - 1.87 1.39 - 1.83 

P-Value 0.123 

Adjusted P-Value 0.265 

Serine 

Mean 1.12 1.12 

0.576 - 2.08 

Range 0.719 - 1.34 0.985 - 1.31 

CI 1.01 - 1.25 1.00 - 1.25 

P-Value 0.971 

Adjusted P-Value 0.971 

Threonine 

Mean 1.11 1.11 

0.619 - 1.95 

Range 0.847 - 1.29 0.997 - 1.26 

CI 1.02 - 1.22 1.01 - 1.21 

P-Value 0.687 

Adjusted P-Value 0.793 

Tryptophan 

Mean 0.325 0.312 

0.153 - 0.537 

Range 0.242 - 0.429 0.236 - 0.427 

CI 0.256 - 0.382 0.239 - 0.371 

P-Value 0.212 

Adjusted P-Value 0.355 

Tyrosine 

Mean 0.635 0.620 

0.337 - 1.20 

Range 0.552 - 0.757 0.508 - 0.737 

CI 0.573 - 0.704 0.560 - 0.688 

P-Value 0.205 

Adjusted P-Value 0.355 

Valine 

Mean 1.39 1.36 

0.682 - 2.49 

Range 1.04 - 1.66 1.20 - 1.59 

CI 1.23 - 1.57 1.20 - 1.54 

P-Value 0.159 

Adjusted P-Value 0.317 a

 The statistical tolerance interval was calculated from commercial canola varieties and calculated to contain with 95 percent 

confidence, 99 percent of the population of canola, negative limits set to zero b

 Least squares mean were generated from a mixed model analysis 

c Range denotes the lowest and highest individual value across sites 

d

 Confidence Interval 

e

 Non-adjusted P-value 

f

 False Discovery Rate (FDR) adjusted P-value
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73496 Canola 

VIII-D. Vitamin Analysis 

Although not specifically mentioned in the OECD consensus document, a standard B vitamin analysis 

was conducted. No statistically significant differences were observed between 73496 and the control 

canola mean values for vitamin B1 (thiamine), vitamin B2 (riboflavin), vitamin B3 (niacin), vitamin B5 

(pantothenic acid), vitamin B6 (pyridoxine), and vitamin B9 (folic acid; Table 18). These results indicate 

73496 canola is comparable to conventional canola with respect to key vitamins. 

Tocopherols are listed in the OECD consensus document as additional important components of canola 

oil as natural antioxidants (OECD, 2001). Concentrations of δ-tocopherol and total tocopherols were 

statistically significantly different between 73496 canola and control canola seed samples using the 

adjusted P-value (Table 18). However, the differences were small in magnitude and, in every case, the 

range of values was within the respective tolerance interval determined using commercial canola 

varieties. The small, statistically significant differences observed for certain tocopherols are unlikely to be 

biologically meaningful. 

In conclusion, vitamin analysis of canola seed demonstrates that 73496 canola is comparable to 

conventional canola. 
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73496 Canola 

Table 18. Summary Analysis of Vitamin Composition in Canola Seed 

Analyte 

Control 

Canola 

n=20 

73496 

Canola 

n=20 

Tolerance 

Intervala

mg/kg Dry Weight

Vitamin B1 

(Thiamine) 

Meanb

 14.2 14.8 

4.92 - 33.1 

Rangec 9.65 - 21.1 10.6 - 25.8 

CId 11.8 - 17.2 12.2 - 17.8 

P-Valuee 0.611 

Adjusted PValuef 0.737 

Vitamin B2 

(Riboflavin) 

Mean 2.70 3.14 

1.45 - 7.33 

Range 2.04 - 3.98 2.27 - 5.88 

CI 2.29 - 3.19 2.66 - 3.71 

P-Value 0.00531 

Adjusted P-Value 0.0507 

Vitamin B3 

(Niacin) 

Mean 188 180 

61.0 - 444 

Range 156 - 245 154 - 208 

CI 179 - 197 171 - 189 

P-Value 0.188 

Adjusted P-Value 0.338 

Vitamin B5 

(Pantothenic Acid) 

Mean 4.74 4.56 

3.52 - 9.88 

Range 4.08 - 5.77 3.94 - 5.05 

CI 4.31 - 5.21 4.15 - 5.01 

P-Value 0.0273 

Adjusted P-Value 0.0974 

Vitamin B6 

(Pyridoxine) 

Mean 3.69 3.39 

1.17 - 20.9 

Range 2.58 - 6.01 2.54 - 5.25 

CI 2.94 - 4.64 2.70 - 4.26 

P-Value 0.0670 

Adjusted P-Value 0.162 

Vitamin B9 

(Folic Acid) 

Mean 2.89 2.63 

0.769 - 5.27 

Range 1.57 - 7.88 1.12 - 5.68 

CI 2.05 - 4.06 1.87 - 3.69 

P-Value 0.307 

Adjusted P-Value 0.458 

α-Tocopherol 

Mean 109 114 

54.7 - 189 

Range 96.5 - 130 101 - 129 

CI 98.8 - 120 103 - 125 

P-Value 0.0179 

Adjusted P-Value 0.0790 

β-Tocopherol 

Mean <1.25g

 <1.25g

0 - 1.67i

Range <1.25g <1.25g

CI NAh NA 

P-Value NA 

Adjusted P-Value NA 

δ-Tocopherol 

Mean 3.23 3.65 

1.93 - 11.9 

Range 2.80 - 4.04 3.25 - 3.99 

CI 3.11 - 3.35 3.51 - 3.78 

P-Value <0.0001 

Adjusted P-Value 0.00216j
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73496 Canola 

Table 18. Summary Analysis of Vitamin Composition in Canola Seed (continued) 

Analyte 

Control 

Canola 

n=20 

73496 

Canola 

n=20 

Tolerance 

Interval 

mg/kg Dry Weight

γ-Tocopherol 

Mean 170 177 

106 - 378 

Range 143 - 196 146 - 205 

CI 155 - 187 161 - 195 

P-Value 0.00770 

Adjusted P-Value 0.0525 

Total Tocopherols 

Mean 283 295 

191 - 499 

Range 243 - 313 250 - 332 

CI 262 - 305 273 - 318 

P-Value 0.00397 

Adjusted P-Value 0.0465j

a

 The statistical tolerance interval was calculated from commercial canola varieties and calculated to contain with 95 

percent confidence, 99 percent of the population of canola, negative limits set to zero b

 Least squares mean were generated from a mixed model analysis 

c Range denotes the lowest and highest individual value across sites 

d

 Confidence Interval 

e

 Non-adjusted P-value 

f

 False Discovery Rate (FDR) adjusted P-value g

 Analyte values were below the assay LLOQ. Sample results below the LLOQ were assigned a value equal to the 

LLOQ. h

 Statistical analysis was not available (NA). 

i Tolerance interval was not available due to insufficient sample values being detected above the assay LLOQ. 

 Minimum and maximum values are used in place of a tolerance interval. 

j Statistically significant difference, FDR adjusted P-Value <0.05. 

VIII-E. Mineral Analysis

Several mineral ions are recognized as essential plant nutrients and are required by the plant in 

significant quantities. These macronutrients include calcium, phosphorus, magnesium, potassium and 

sodium. The micronutrient minerals, iron, copper, manganese and zinc are incorporated in plant tissues 

in only trace amounts. Both macro- and micro-nutrient minerals were analyzed in seed samples from 

73496 canola and compared with corresponding values from samples of near-isoline control canola seed. 

A statistically significant difference was observed in magnesium concentration between 73496 and control 

canola using the adjusted P-value (Table 19). The magnitude of the difference was small and the range 

of individual values was within the tolerance interval determined for seed magnesium concentrations 

using commercial canola varieties. 

In conclusion, mineral analysis of canola seed demonstrates that 73496 canola is comparable to 

conventional canola. 
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73496 Canola 

Table 19. Summary Analysis of Mineral Composition in Canola Seed 

Analyte Control Canola

n=20 

73496 Canola

n=20 

Tolerance

Intervala

% Dry Weight 

Calcium 

Meanb

 0.465 0.474 

0.172 - 0.939 

Rangec 0.365 - 0.576 0.397 - 0.572 

CId 0.406 - 0.532 0.414 - 0.542 

P-Valuee 0.457 

Adjusted P-Valuef 0.625 

Phosphorus 

Mean 0.788 0.756 

0.299 - 1.58 

Range 0.543 - 1.13 0.567 - 1.14 

CI 0.608 - 1.02 0.583 - 0.980 

P-Value 0.0285 

Adjusted P-Value 0.0974 

Magnesium 

Mean 0.357 0.325 

0.219 - 0.538 

Range 0.308 - 0.410 0.277 - 0.385 

CI 0.327 - 0.390 0.298 - 0.356 

P-Value <0.0001 

Adjusted P-Value 0.00216g

Manganese 

Mean 0.00381 0.00351 

0.00174 - 

0.00769 

Range 0.00275 - 0.00463 0.00278 - 0.00418 

CI 0.00318 - 0.00458 0.00292 - 0.00422 

P-Value 0.0334 

Adjusted P-Value 0.110 

Copper 

Mean 0.000175 0.000185 

0 - 0.000772 

Range <0.000125h - 0.000251 <0.000125h - 0.000276 

CI 0.000129 - 0.000236 0.000137 - 0.000250 

P-Value 0.0972 

Adjusted P-Value 0.221 

Iron 

Mean 0.00677 0.00634 

0.00163 - 

0.0259 

Range 0.00452 - 0.0200 0.00412 - 0.0116 

CI 0.00427 - 0.0107 0.00400 - 0.0100 

P-Value 0.288 

Adjusted P-Value 0.454 

Potassium 

Mean 0.649 0.659 

0.284 - 1.79 

Range 0.489 - 0.866 0.487 - 0.898 

CI 0.525 - 0.803 0.533 - 0.816 

P-Value 0.401 

Adjusted P-Value 0.567 

Sodium 

Mean 0.00168 0.00179 

0.0000886 - 

0.0270 

Range 0.000568 - 0.00599 0.000849 - 0.00505 

CI 0.000790 - 0.00358 0.000840 - 0.00381 

P-Value 0.507 

Adjusted P-Value 0.671 

Zinc 

Mean 0.00398 0.00384 

0.00163 - 

0.0101 

Range 0.00277 - 0.00548 0.00233 - 0.00594 

CI 0.00317 - 0.00498 0.00306 - 0.00481 

P-Value 0.288 

Adjusted P-Value 0.454 a

 The statistical tolerance interval was calculated from commercial canola varieties and calculated to contain with 95 percent

confidence, 99 percent of the population of canola, negative limits set to zero b

 Least squares mean were generated from a mixed model analysis 

c

 Range denotes the lowest and highest individual value across sites 

d

 Confidence Interval 

e

 Non-adjusted P-value 

f

 False Discovery Rate (FDR) adjusted P-value 

g

 Statistically significant difference, FDR adjusted P-Value <0.05. h

 Analyte values were below the assay LLOQ. Sample results below the LLOQ were assigned a value equal to the LLOQ.
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VIII-F. Glucosinolate Analysis

Glucosinolates are considered key toxicants of canola (OECD, 2001). The major glucosinolates in canola 

are 3-butenyl glucosinolate (gluconapin), 4-pentenyl glucosinolate (glucobrassicanapin), 2- hydroxy-3-

butenyl glucosinolate (progoitrin) and 2-hydroxy-4-pentenyl glucosinolate (napoleiferin). 

Glucosinolates themselves are generally considered to be innocuous; however the hydrolysis products 

have negative effects on animal production. The low palatability and the adverse effects of glucosinolates 

due to their antithyroid activity led to the development of varieties of rapeseed which have combined low 

concentrations of both glucosinolates and erucic acid (also known as “double low” varieties). The total 

glucosinolate concentration for 73496 canola was 5.66 μmoles/g dry weight, which is within the 

acceptable definition for canola (maximum 30 μmoles/g dry weight; OECD, 2001). 

A statistically significant difference was observed in progoitrin concentration between 73496 canola and 

control canola samples based on the adjusted P-value. The range of concentrations of progoitrin in 73496 

canola was within the tolerance intervals established using commercial canola varieties (Table 20). 

In conclusion, glucosinolate analysis of canola seed demonstrates that 73496 canola is comparable to 

conventional canola. 
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73496 Canola 

Table 20. Summary Analysis of Glucosinolate Composition in Canola Seed 

Analyte Control Canola 

n=20 

73496 Canola 

n=20 

Tolerance

Intervala

μmol/g Dry Weight

Glucoiberin 

Meanb

 <0.176g <0.176g

NCi

Rangec <0.176g <0.176g

CId NAh NA 

P-Valuee NA 

Adjusted PValuef NA 

Progoitrin 

Mean 0.524 0.412 

0.130 - 11.6 

Range 0.224 - 1.29 0.181 - 1.37 

CI 0.233 - 1.18 0.183 - 0.928 

P-Value 0.00174 

Adjusted P-Value 0.0238j

Epi-Progoitrin 

Mean 0.0717 <0.0708g

0 - 0.159k

Range <0.0708g - 0.0883 <0.0708g

CI NA NA 

P-Value NA 

Adjusted P-Value NA 

Glucoraphanin 

Mean 0.0847 0.0789 

0 - 0.565 

Range <0.0488g - 0.217 <0.0488g - 0.143 

CI 0.0655 - 0.110 0.0610 - 0.102 

P-Value 0.401 

Adjusted P-Value 0.567 

Gluconapoleiferin 

Mean 0.0339 0.0330 

0 - 0.372 

Range <0.0299g - 0.0618 <0.0299g - 0.0544 

CI 0.0280 - 0.0411 0.0272 - 0.0400 

P-Value 0.272 

Adjusted P-Value 0.446 

Gluconapin 

Mean 0.759 0.773 

0.197 - 10.0 

Range 0.286 - 2.01 0.304 - 2.66 

CI 0.328 - 1.75 0.335 - 1.79 

P-Value 0.777 

Adjusted P-Value 0.868 

Glucoalyssin 

Mean 0.0753 0.0897 

0 - 0.663 

Range <0.0676g - 0.125 <0.0676g - 0.223 

CI 0.0508 - 0.111 0.0606 - 0.133 

P-Value 0.0496 

Adjusted P-Value 0.133 

4-Hydroxyglucobrassicin 

Mean 3.36 3.41 

1.28 - 11.3 

Range 1.53 - 6.06 1.78 - 8.33 

CI 2.40 - 4.71 2.44 - 4.76 

P-Value 0.836 

Adjusted P-Value 0.911 

Glucobrassicin 

Mean 0.404 0.383 

0.0376 - 

5.25 

Range 0.215 - 0.787 0.206 - 1.14 

CI 0.271 - 0.600 0.257 - 0.569 

P-Value 0.189 

Adjusted P-Value 0.338 
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73496 Canola 

Table 20. Summary Analysis of Glucosinolate Composition in Canola Seed (continued) 

Analyte Control Canola 

n=20 

73496 Canola 

n=20 

Tolerance

Interval 

μmol/g Dry Weight

Glucobrassicinapin 

Mean 0.401 0.395 

0 - 1.80 

Range <0.390g - 0.475 <0.390g - 0.441 

CI NA NA 

P-Value NA 

Adjusted P-Value NA 

Gluconasturtiin 

Mean 0.255 0.286 

0 - 5.39 

Range 0.133 - 0.991 0.139 - 0.610 

CI 0.166 - 0.391 0.186 - 0.439 

P-Value 0.302 

Adjusted P-Value 0.458 

4-Methoxyglucobrassicin 

Mean <0.0774g

 <0.0774g

NC 

Range <0.0774g <0.0774g

CI NA NA 

P-Value NA 

Adjusted P-Value NA 

Neoglucobrassicin 

Mean 0.0466 0.0641 

0 - 0.192 

Range <0.0198g - 0.0903 0.0230 - 0.162 

CI 0.0266 - 0.0815 0.0366 - 0.112 

P-Value 0.00897 

Adjusted P-Value 0.0525 

Total Glucosinolates 

Mean 5.77 5.66 

2.17 - 30.0 

Range 2.61 - 10.5 2.77 - 13.5 

CI 3.72 - 8.94 3.65 - 8.77 

P-Value 0.686 

Adjusted P-Value 0.793 a

 The statistical tolerance interval was calculated from commercial canola varieties and calculated to contain with 95 

 percent confidence, 99 percent of the population of canola, negative limits set to zero b

 Least squares mean were generated from a mixed model analysis 

c Range denotes the lowest and highest individual value across sites 

d

 Confidence Interval 

e Non-adjusted P-value f False Discovery Rate (FDR) adjusted P-value g Theoretical LLOQ values were calculated by mathematically proportioning a known or measured canola seed 

glucosinolate concentration per measured signal-to-noise (S:N) ratio to a theoretical glucosinolate concentration 

per S:N of 10. Sample results below the theoretical LLOQ were designated < [LLOQ]. h Statistical analysis was not available (NA). i Tolerance interval could not be calculated (NC) due to all sample values below the assay LLOQ. j Statistically significant difference, FDR adjusted P-Value <0.05. k Tolerance interval was not available due to insufficient sample values being detected above the assay LLOQ. 

Minimum and maximum values are used in place of a tolerance interval. 
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VIII-G. Secondary Metabolite and Anti-Nutrient Analysis

Tannins, sinapine, and phytic acid are considered to be anti-nutrients in canola meal (OECD, 2001). A 

major phenolic compound in canola, sinapine imparts a bitter taste to canola meal (OECD, 2001). Phytic 

acid is the major form of phosphorus in plants; however, it is unavailable as a nutrient source for animals 

(OECD, 2001). 

Phytosterols are cholesterol-like molecules found in all plant foods, with the highest concentrations 

occurring in vegetable oils. They are absorbed only in trace amounts but have the beneficial effect of 

inhibiting the absorption of dietary cholesterol (Ostlund, 2002). Phytosterols are not endogenously 

synthesized in the body but are derived solely from the diet (Rao and Koratkar, 1997). 

The only statistically significant difference found between 73496 compared with control canola was for 

cholesterol based on the adjusted P-value. However, the range of values for cholesterol was within the 

established tolerance interval and is unlikely to be biologically meaningful (Table 21). 

In conclusion, secondary metabolite and anti-nutrient analysis of canola seed demonstrates that 73496 

canola is comparable to conventional canola. 
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73496 Canola 

Table 21. Summary Analysis of Secondary Metabolite and Anti-Nutrient Composition in Canola 

Seed 

Analyte Control Canola 

n=20 

73496 Canola 

n=20 

Tolerance

Intervala

% Dry Weight

Tannins-Soluble 

Meanb 0.113 0.109 

0.0521 - 

0.297 

Rangec 0.0768 - 0.180 0.0777 - 0.176 

CId 0.0840 - 0.152 0.0812 - 0.147 

P-Valuee 0.415 

Adjusted P-Valuef 0.577 

Tannins-Insoluble 

Mean 0.330 0.237 

0.0731 - 

2.32 

Range 0.155 - 0.824 0.142 - 0.492 

CI 0.218 - 0.499 0.157 - 0.359 

P-Value 0.00874 

Adjusted P-Value 0.0525 

Phytic Acid 

Mean 1.84 1.73 

0.684 - 6.06 

Range 1.04 - 2.86 1.01 - 3.00 

CI 1.27 - 2.68 1.19 - 2.52 

P-Value 0.0401 

Adjusted P-Value 0.126 

Sinapine 

Mean 0.928 0.937 

0.538 - 1.33 

Range 0.763 - 1.12 0.804 - 1.09 

CI 0.852 - 1.01 0.861 - 1.02 

P-Value 0.558 

Adjusted P-Value 0.727 

Cholesterol 

Mean 0.000866 0.00110 

0 - 0.00238 

Range 0.000685 - 0.00121 0.000850 - 0.00135 

CI 0.000801 - 0.000938 0.00101 - 0.00119 

P-Value 0.000281 

Adjusted P-Value 0.00461g

Brassicasterol 

Mean 0.0147 0.0160 

0.00830 - 

0.108 

Range 0.0106 - 0.0190 0.0124 - 0.0200 

CI 0.0129 - 0.0168 0.0141 - 0.0183 

P-Value 0.0230 

Adjusted P-Value 0.0945 

Campesterol 

Mean 0.0921 0.0990 

0.0249 - 

0.257 

Range 0.0738 - 0.111 0.0707 - 0.125 

CI 0.0775 - 0.109 0.0833 - 0.118 

P-Value 0.0492 

Adjusted P-Value 0.133 

Stigmasterol 

Mean 0.00269 0.00268 

0.000832 - 

0.0113 

Range 0.00234 - 0.00316 0.00228 - 0.00335 

CI 0.00248 - 0.00292 0.00247 - 0.00291 

P-Value 0.942 

Adjusted P-Value 0.957 

β-Sitosterol 

Mean 0.132 0.137 

0.0428 - 

0.387 

Range 0.0967 - 0.158 0.103 - 0.174 

CI 0.115 - 0.151 0.120 - 0.157 

P-Value 0.176 

Adjusted P-Value 0.328 
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73496 Canola 

Table 21. Summary Analysis of Secondary Metabolite and Anti-Nutrient Composition in Canola 

Seed (continued) 

Analyte Control Canola 

n=20 

73496 Canola 

n=20 

Tolerance

Interval 

% Dry Weight

Total Sterols 

Mean 0.242 0.257 

0.0801 - 

0.741 

Range 0.189 - 0.287 0.206 - 0.323 

CI 0.215 - 0.274 0.228 - 0.290 

P-Value 0.0846 

Adjusted P-Value 0.198 a

 The statistical tolerance interval was calculated from commercial canola varieties and calculated to contain with 95 

percent confidence, 99 percent of the population of canola, negative limits set to zero b

 Least squares mean were generated from a mixed model analysis 

c Range denotes the lowest and highest individual value across sites 

d

 Confidence Interval 

e

 Non-adjusted P-value 

f False Discovery Rate (FDR) adjusted P-value g

 Statistically significant difference, FDR adjusted P-Value <0.05. 

VIII-H. Acetylated Amino Acids 

The GAT4621 enzyme exhibits measureable activity with five amino acid substrates (L-aspartate, 

L-glutamate, L-serine, glycine and L-threonine) in addition to glyphosate (Appendix 7). In studies using 

microbial-produced GAT4621, the level of catalytic efficiency of GAT4621 on aspartate, glutamate, 

serine, and threonine was 1%, 0.8%, 0.05%, and 0.06%, respectively, of that observed for glyphosate 

(Pioneer data not shown). The affinity of the GAT4621 enzyme for glycine was too low to estimate the 

level of catalytic efficiency. Levels of activity with other tested substrates, including a wide range of other 

amino acids and antibiotics, were below the limit of quantification (Pioneer data not shown). 

Based on the activity of the GAT4621 enzyme, the concentrations of N-acetylaspartate (NAA), 

N-acetylglutamate (NAG), N-acetylserine (NAS), N-acetylthreonine (NAT), and N-acetylglycine (NAGly) 

were measured in samples of seed, whole plant, and processed fractions derived from 73496 canola and 

control canola. 

H1. Acetylated Amino Acids in Seed and Whole Plant Samples 

Concentrations of the five acetylated amino acids were measured in seed samples of 73496 canola and 

control canola. Results are provided in Table 22 below. As expected based on catalytic efficiency, NAA 

and NAG were the two most abundant in 73496 canola. Mean concentrations of NAA and NAG were 

1480 μg/g dry weight and 32.8 μg/g dry weight, respectively, for 73496 canola seed samples (Table 22). 

Although the mean concentrations of NAT and NAS were elevated (statistically significant) in 73496 

canola seed, relative to the near-isoline control line, the range of individual values for both these 

acetylated amino acids were within the tolerance interval established using commercial canola varieties 

(Table 22). There was no statistically significant difference in NAGly concentration measured in 73496 

canola seed samples relative to the control line. 
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73496 Canola 

Table 22. Concentrations of Acetylated Amino Acids in Canola Seed 

Analyte 

Control Canola 

n=20 

73496 Canola 

Treated 

n=20 

Tolerance 

Intervala

μg/g Dry Weight 

NAA 

Meanb

 1.24 1480 

0.00861 - 4.43 

Rangec 0.377 - 5.39 1200 - 1770 

CId 0g - 9.38 1340 - 1640 

P-valuee <0.0001 

Adjusted P-valuef <0.0001g

NAG 

Mean 0.628 32.8 

0.0968 - 5.37 

Range 0.428 - 1.46 20.3 - 61.1 

CI 0.00000752 - 2.50 24.4 - 42.5 

P-value <0.0001 

Adjusted P-value <0.0001h

NAGly 

Mean 0.0751 0.0825 

0.0240 - 0.338 

Range 0.0481 - 0.125 0.0424 - 0.182 

CI 0.0540 - 0.105 0.0592 - 0.115 

P-value 0.454 

Adjusted P-value 0.454 

NAS 

Mean 0.843 1.04 

0.0524 - 27.2 

Range 0.389 - 3.05 0.491 - 3.55 

CI 0.437 - 1.63 0.542 - 2.01 

P-value 0.0035 

Adjusted P-value 0.00528h

NAT 

Mean 0.110 0.546 

0.0140 - 1.74 

Range 0.0531 - 0.212 0.260 - 1.64 

CI 0.0665 - 0.181 0.331 - 0.902 

P-value <0.0001 

Adjusted P-value <0.0001h

a

 The statistical tolerance interval was calculated from commercial canola varieties and calculated to contain with 95 

percent confidence, 99 percent of the population of canola, negative limits set to zero b

 Least squares mean were generated from a mixed model analysis 

c Range denotes the lowest and highest individual value across sites 

d

 Confidence Interval 

e

 Non-adjusted P-value 

f False Discovery Rate (FDR) adjusted P-value g

 The lower limit of the confidence interval was negative on the transformed scale and was set to zero.prior to backtransformation 

h

 Statistically significant difference, FDR adjusted P-Value <0.05. 
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73496 Canola 

Concentrations of the five acetylated amino acids were measured in whole plant samples derived from 

greenhouse-grown 73496 canola and control canola. Concentrations of all measured acetylated amino 

acids except NAS were elevated in whole plant samples of 73496 canola plants compared to control 

(Table 23). Consistent with results obtained for seed samples obtained from 73496 canola, NAA and 

NAG were the two most abundant with mean concentrations of 4560 μg/g and 26 μg/g, in whole plant 

samples, respectively. 

Table 23. Concentrations of Acetylated Amino Acids in Canola Whole Plant Samples 

Analyte 

Control 

Canola 

n=15 

73496 

Canola 

n=15 

μg/ga

NAA 

Meanb

 0.705 4560 

Rangec 0.404 - 1.23 3730 - 5340 

CId 0.649 - 0.766 4190 - 4950 

P-valuee

 <0.0001 

Adjusted P-valuef <0.0001g

NAG 

Mean 2.04 26.0 

Range 1.45 - 3.27 21.0 - 35.9 

CI 1.87 - 2.22 23.8 - 28.3 

P-value <0.0001 

Adjusted P-value <0.0001g

NAGly 

Mean 0.152 0.344 

Range 0.122 - 0.193 0.247 - 0.445 

CI 0.139 - 0.166 0.316 - 0.376 

P-value <0.0001 

Adjusted P-value <0.0001g

NAS 

Mean 14.0 13.0 

Range 10.2 - 21.9 9.17 - 22.0 

CI 12.3 - 15.9 11.5 - 14.7 

P-value 0.406 

Adjusted P-value 0.406 

NAT 

Mean 1.92 7.67 

Range 1.60 - 2.50 5.43 - 11.8 

CI 1.76 - 2.11 7.00 - 8.39 

P-value <0.0001 

Adjusted P-value <0.0001g

a

 Results reported as-is after lyophilization without adjusting for moisture b

 Least squares mean were generated from a mixed model analysis 

c Range denotes the lowest and highest individual value across sites. 

d

 Confidence Interval 

e

 Non-adjusted P-value 

f False Discovery Rate (FDR) adjusted P-value 

g

 Statistically significant difference, FDR adjusted P-Value <0.05. 
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H2. Acetylated Amino Acids in Processed Fractions 

Whole canola seed is not commonly consumed. Because the typical processed canola products 

consumed by animals and humans are meal and oil, respectively, the concentrations of the five

acetylated amino acids were measured in processed fractions of 73496 canola and control canola. 

As expected, based on data from whole seed samples, NAA and NAG were the two most abundant 

acetylated amino acids in 73496 canola meal. Mean concentrations of NAA and NAG were 2872 μg/g 

and 62.3 μg/g, respectively, for 73496 canola toasted meal from seed with hulls (i.e., seed coats) (Table 

24). The corresponding concentrations from toasted meal produced from canola seed without hulls of 

each event were similar at 3013 and 66.0 μg/g, respectively for NAA and NAG. The concentrations of 

each of the remaining acetylated amino acids, NAT, NAS and NAGly, were approximately 30–100 fold 

less than the concentration of NAG measured in toasted meal produced from 73496 canola seed either 

with or without hulls. 

On an equivalent weight basis, the elevated concentrations of acetylated amino acids in toasted meal, 

relative to concentrations measured in whole seed samples, reflect removal of the oil component, which is 

not expected to contain these compounds. This was confirmed through analysis of oil samples, where 

the concentrations of each acetylated amino acid were either not detectable or below the limit of 

quantification in all samples of refined, bleached, deodorized oil (Table 24). 

Table 24. Concentrations of Acetylated Amino Acids in Processed Fractions Produced from 73496 

Canola and Control Canola 

Fraction Canola 

Line

Concentration 

(Range) 

n=2a 

NAAb NAG NAT NAS NAGly 

μg/gc

Toasted Meal 

(with hulls) 

73496 2872 

(2822 - 2921) 

62.3 

(57.7 - 66.8) 

0.585 

(0.558 - 0.612) 

1.20 

(1.11 - 1.29) 

0.151 

(0.145 - 0.157) 

Control 2.31 

(2.15 - 2.47) 

1.36 

(1.31 - 1.41) 

0.161 

(0.146 - 0.175) 

1.46 

(1.39 - 1.53) 

0.129 

(0.114 - 0.144) 

Toasted Meal 

(without hulls) 

73496 3013 

(3002 - 3024) 

66.0 

(62.6 - 69.3) 

0.788 

(0.535 - 1.04) 

2.27 

(1.98 - 2.55) 

0.224 

(0.184 - 0.265) 

Control 2.58 

(2.00 - 3.16) 

1.71 

(1.63 - 1.80) 

0.226 

(0.195 - 0.258) 

1.76 

(1.51 - 2.01) 

0.180 

0.163 - 0.197) 

RBD Oil 

(with hulls) 

73496 NDd

 ND - <LLOQ ND - <LLOQ ND ND - <LLOQ 

Control ND ND <LLOQ ND ND - <LLOQ 

RBD Oil 

(without hulls) 

73496 ND - <LLOQe

 <LLOQ ND ND ND -<LLOQ 

Control ND ND ND ND ND 

a

 Statistics are not provided due to small sample size. 

b

 NAA = N-acetylaspartate; NAG = N-acetylglutamate; NAT = N-acetylthreonine; NAS = N-acetylserine; NAGly = Nacetylglycine c Values are not corrected for moisture content. d

 ND = Not detected 

e

 LLOQ = Lower limit of quantification, which was 0.01 μg/g for a 100 mg sample 

VIII-I. Conclusions on the Compositional Analysis of 73496 Canola

The analytes for compositional assessment were selected considering the OECD consensus document 

on compositional considerations for new varieties of canola (OECD, 2001). Among the numerous 
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73496 Canola 

compositional analyses that were carried out, concentrations of most analytes were not significantly 

different between 73946 canola and control canola. Statistically significant differences were noted for 

concentrations of oleic and linoleic fatty acids; delta- and total tocopherols; magnesium; the glucosinolate 

progoitrin; and cholesterol. However, the magnitudes of the differences were small and in every case the 

ranges of values was all within the respective tolerance interval established using commercial canola 

varieties. Overall, no consistent patterns emerged to suggest that biologically significant changes in 

composition or nutritive value of the seed had occurred as an unexpected result of the transformation 

process. 

In addition to the compositional analytes per OECD guidelines, the concentrations of NAA, NAG, NAS, 

NAT, and NAGly were measured in samples of seed, whole plant, and processed fractions derived from 

73496 canola. As described earlier, these analyses were conducted because the GAT4621 protein is 

known to acetylate certain free amino acids (L-aspartate, L-glutamate, glycine, L-serine, and L-threonine) 

resulting in the production of NAA, NAG, NAGly, NAS, and NAT, respectively. As expected, NAA and 

NAG were the most abundant in seed, whole plant, and processed meal fractions. Refined, bleached, 

deodorized oil fractions did not contain measurable levels of these five acetylated amino acids. These 

acetylated amino acids are normal components of food and feed, have a safe history of food and feed 

use, and are not novel substances (Appendix 8). The safety of these substances has also been 

evaluated in published studies as described in Appendix 8; therefore, it is not expected that the increase 

in these acetylated amino acids would adversely affect the safety of processed products from 73496 

canola. 

Based on the OECD guidelines for compositional equivalence, we have concluded that 73496 canola was 

compositionally comparable to conventional canola. The increases in certain acetylated amino acids do 

not negatively impact the safety of 73496 canola and will not have an adverse impact on processed 

commodity products from 73496 canola. 
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73496 Canola 

IX. Assessment of Plant Pest Risk for 73496 Canola 

In this section, the plant pest risk of 73496 canola is discussed with respect to the potential environmental 

impact of the presence of the GAT4621 protein in 73496 canola, the fate of the gat4621 gene in humans 

and animals, the weediness potential of 73496 canola, and the impact of gene flow from 73496 canola to 

crop relatives or wild relatives of canola. 

IX-A. Environmental Impact of the Transgenic Protein 

The GAT4621 protein present in 73496 canola is identical to the GAT4621 protein present in previously 

authorized maize event DP-Ø9184Ø-6 (FDA, 2008; USDA, 2009). The GAT4621 protein was reviewed 

by FDA in a New Protein Consultation submitted to US FDA on January 31, 2007 and completed on 

October 7, 2009. 

The GAT4621 protein is unlikely to pose a hazard to the environment. The GAT4621 protein sequence is 

derived from N-acetyltransferase protein sequences from Bacillus licheniformis, a gram positive 

saprophytic bacterium that is ubiquitous in soil. The GAT4621 protein is a member of the GNAT 

acetyltransferase superfamily that contains more than 10,000 representatives from plants, animals, 

bacteria, and fungi, all of which share a highly conserved GNAT motif (Section VI) (Vetting et al., 2005). 

GAT4621 is 75-78% identical and 90-91% similar at the amino acid level to the translated protein 

sequences of each of the three original gat genes from Bacillus licheniformis from which the gat4621

gene was derived. GAT4621 retains the acetyltransferase enzyme function of the native proteins. In 

addition, the GAT4621 protein is highly unlikely to be a toxin (Section VI). 

Based on the facts above, there are not likely to be any adverse environmental effects due to the 

presence of the GAT4621 protein in 73496 canola. 

IX-B. Fate of Transgenic DNA in Humans and Animals 

Transgenic DNA is no different from other DNA consumed as part of the normal diet. Genetically 

engineered organisms have been used in drug production (Thayer, 2005) and microbial fermentation 

(cheese) since the late 1970’s (Maryanski, 1995; National Centre for Biotechnology Education, 2006). 

More than 2.47 billion cumulative acres (1 billion hectares) of engineered food and feed crops have been 

grown and consumed worldwide from 1996 to 2010 (ISAAA, 2010). The FDA has not reported any 

significant concerns with bioengineered food and feed currently on the market. The EPA has exempted 

from a tolerance DNA that encodes currently registered plant incorporated protectants because of a lack 

of toxicity (Federal Register, 2001). 

Studies in humans and animals following the fate of DNA once consumed have shown that the majority of 

DNA is degraded in the gastrointestinal tract. There is evidence that DNA can move from the 

gastrointestinal tract lumen to other areas of the body, but this is considered to be a normal occurrence 

and no risks have been identified as a result of absorption (Duggan et al., 2003; Einspanier et al., 2001). 

In conclusion, there are not likely to be any adverse effects due to the presence of the gat4621 gene and 

associated transgenic DNA in 73496 canola. 

IX-C. Weediness Potential of 73496 Canola 

Canola is not considered a noxious or invasive weed in the United States (USDA-NRCS, 2010). Weed 

species typically spread easily in disturbed areas or within crops and generally have a range of life history 

characteristics in common that enables them to rapidly colonize and persist in an ecosystem. Several 

characteristics of weeds have been described by Baker (1974) and are indicated in Table 25. In general, 

these characteristics do not apply to cultivated canola varieties. 
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Table 25. Common Characteristics of Weeds 

Characteristic Example Applies to canola varieties? 

Dissemination of seed Long continuous seed 

production 

No. As a cultivated crop, canola has 

been selectively bred to ensure 

seed production is timely to allow for 

efficient harvest. Additionally, canola 

is an early successional ruderal (i.e., 

capable of growing in disturbed 

habitats) which is incapable of 

regenerating in undisturbed habitats 

(Crawley et al., 1993). Populations 

were observed to be seed limited in 

a study conducted on motorway 

verges by Crawley and Brown 

(1995). 

Special adaptations for short 

and long-distance dispersal 

Possible. However, the small size of 

canola seeds and their high 

numbers on post-harvest fields may 

facilitate some wind dispersal 

(Lutman, 1993). The dispersal 

distance will depend on wind 

strength, the amount of debris on 

the ground and the moisture content 

of the material. It is reasonable to 

expect that seeds and pods of low 

moisture content may be 

transported within the field to 

adjacent fields, or outside 

agricultural areas. 

High seed output under 

favorable conditions 

Yes. However due to selective 

breeding, crop plants only functional 

optimally under managed 

agricultural conditions, such as high 

soil fertility or low plant competition. 

These conditions rarely occur in 

natural habitats, resulting in poor 

fitness of canola plants (i.e. reduced 

recruitment, low survivorship, poor 

competitive ability, low seed 

production) (OGTR, 2002). 

Dormancy of seed Continuous germination and 

long-lived seeds 

Yes. Studies in the Northern 

Hemisphere have reported viable 

seeds of canola persisting in 

disturbed soils for at least 5 years 

and possibly up to 10 years or more 

in undisturbed soil (Chadoeuf et al., 

1998; Madsen 1962; Pekrun et al., 

1997; Vaughan et al., 1976). 
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Table 25. Common Characteristics of Weeds (continued) 

Characteristic Example Applies to canola varieties? 

Germination of seed/seed 

survival 

Germination and seed 

production under a wide 

range of environmental 

conditions 

Possible. The persistence of canola 

seeds in a seed bank depends both 

on seed dormancy and their vertical 

distribution in the soil, as seeds are 

more likely to persist at deep rather 

than shallow depths (Pekrun et al., 

1998; Simard et al., 2002). 

As with many annual weeds, feral 

canola plants usually are not 

capable of surviving outside of 

cultivation and without human 

intervention for more than a few 

generations (Andersson and de 

Vicente, 2010). Still, some old 

canola cultivars can persist outside 

of cultivation for up to eight or nine 

years after they were last cultivated, 

albeit at low densities (Gulden et al., 

2003; Légère et al., 2001; Simard et 

al., 2002; Squire et al., 1999). 

Competitiveness Choking growth or production 

of allelochemicals 

No. Canola is not known to produce 

allelochemicals or otherwise 

strongly compete with other plants. 

Canola is known to be strongly 

affected by weed competition during 

crop establishment (Berglund et al., 

2007) and is unable to invade 

established natural habitats (OGTR, 

2002). 

Ability to regenerate from 

severed rootstock 

No. Canola is not known to 

regenerate from severed rootstock. 

Rapid seedling growth No. Young canola seedlings are 

very sensitive to early weed 

competition (Berglund et al., 2007). 

Canola is a poor competitor and not 

regarded as an environmentally 

hazardous colonizing species. 

Unless the habitat is regularly 

disturbed, or seed replenished from 

outside, canola will be displaced by 

other plants (OGTR, 2002). 

Rapid growth to reproductive 

stage 

No. As a crop, canola has been 

selectively bred to develop to 

maturity for the purpose full yield 

potential. This is in contrast to rapid 

growth to reproductive stage for the 

purpose of dispersing pollen, as with 

weed species. 
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Table 25. Common Characteristics of Weeds (continued) 

Characteristic Example Applies to canola varieties? 

 Self-compatible, but not 

obligatorily self-pollinated or 

apomictic 

Yes. Fertilization of ovules usually 

results from self pollination, 

although out-crossing rates of 20-

30% have been reported (Rakow 

and Woods, 1987). The floral 

biology of canola is such that it is 

suited to cross pollination by insect 

vectors, but can be pollinated by 

wind (Cresswell et al., 2004; Hayter 

and Cresswell, 2006; Hoyle et al., 

2007). 

Stress tolerance High tolerance or plasticity of 

climatic and edaphic variation 

Possible. Canola grows well in dry 

environments and can tolerate 

moderately saline soil conditions 

(Bañuelos et al., 1997; Stricker et 

al., 1997). However, although 

tolerant of some drought stress, 

seed yields of Brassica napus are 

known to decrease due to drought 

stress (Jensen et al., 1996; Kumar 

and Singh, 1998) and are most 

susceptible to heat and drought 

stress during flowering (Berglund et 

al., 2007). 

It is generally accepted that most crop plants, including canola, have undergone many years of selective 

breeding and domestication and therefore, they function optimally only under managed agricultural 

conditions. Similar conditions rarely occur in natural habitats, resulting in poor fitness of canola plants 

outside of a managed field (OGTR, 2002). Although canola has a number of life history traits in common 

with those usually associated with weeds (Table 25), it is a poor competitor and is not regarded as a 

colonizing species (Salisbury, 2002). Unless the habitat is regularly disturbed or seeds are replenished 

from outside, canola will be displaced by other plants (Salisbury, 2002). 

The agronomic characteristics and germination data (Section VII) provide evidence that the genetic 

modification resulting in 73496 canola did not alter any major characteristics of the plant that would allow 

for development of weedy characteristics different from other canola varieties. Furthermore, the 

herbicide-tolerance trait conferred by gat4621 gene does not provide a selective advantage in 

unmanaged ecosystems, but rather only in settings where glyphosate is being applied for weed control. 

73496 canola is susceptible to other herbicides, so control of volunteers could be achieved by use of 

alternative herbicides, or by non-chemical methods that may be part of the weed management best 

practices. 

There is no evidence that herbicide-tolerant varieties of canola are more invasive or more persistent in 

disturbed habitats than their unmodified counterparts, nor does canola survive well in the wild (Crawley et 

al., 1993; Crawley et al., 2001), therefore it is highly unlikely to invade other habitats (Salisbury, 2002). 

Additionally, because there are many different traits that lead to weediness in plants (e.g., those 

described in Table 25), one can assume that these are likely to be polygenic traits and changes to these 

traits would not be easily conferred by adding a single herbicide tolerance gene. 

Herbicide-tolerant canola, including varieties tolerant to glyphosate, has been cultivated since the 

mid-90’s (Devine, 2005). To our knowledge, there is no record of herbicide-tolerant canola varieties 
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demonstrating weedy characteristics as a result of their herbicide tolerance characteristics. Based on 

Pioneer’s detailed characterization, we expect 73496 canola to be similar to other herbicide-tolerant 

canola with regard to weediness characteristics. 

IX-D. Gene Flow Assessment 

Gene flow is the movement of genes via pollen flow or seed dispersal; pollen-mediated gene flow could 

be a potential route for transfer of traits from canola to other related crops or to wild relatives. The 

introgression of a gene (e.g., the introduced gat4621 gene) into another species is limited by several 

variables including pollen viability and dispersal, synchrony of flowering or pollen production, wind speed 

and direction, topography and surrounding vegetation, temperature, humidity, relative density of donor 

and receptor plant populations, sexual incompatibility, and genetic instability (Eastham and Sweet, 2002; 

Gliddon et al., 1999; Ingram, 2000; Thompson et al., 1999; Warwick et al., 2009). In order to evaluate the 

potential for gene flow from 73496 canola, the geographic areas of canola production, the crop relatives 

of canola, the wild relatives of canola, and the barriers to gene flow are discussed further below. In order 

to be conservative in the analysis of possible gene flow, crop or wild relative species were considered for 

further examination if the species had geographical prevalence in canola production areas and 

hybridization with B. napus was possible. 

D1. Geographic Areas of Canola Production 

Canola is grown in the U.S. and the primary states of cultivation include North Dakota (approximately 

90% of total production), Minnesota, Idaho, Washington, Montana, Oklahoma, and Oregon (USDA-ERS, 

2010; USDA-NASS, 2009; USDA-NASS, 2010). Spring and winter canola are generally grown in different 

regions of the U.S. based on climate zones most suitable for the varieties (Mills, 1996; Figure 19). Spring 

canola (typically planted in March and harvested around September to October) in the U.S. is primarily 

grown in areas of North Dakota (90% of production acreage), Minnesota, and the Pacific Northwest (e.g. 

Idaho, Washington, and Montana), due to spring canola’s poor performance in warm temperatures. 

Winter canola (typically planted in September and harvested around June to July) in the U.S. is grown on 

fewer acres than spring canola but can be grown in a broader range of environments if winters are mild. 

In recent years, winter canola variety trial programs are run in the Southeast (Alabama, Arkansas, 

Virginia, Georgia), the Midwest (Illinois, Kentucky, Michigan, Ohio, Pennsylvania), and the Great Plains 

(Wyoming, Colorado, Nebraska, Kansas, Oklahoma, Texas, Missouri); these states also have reported 

canola production (Brown et al., 2008; USDA-NASS, 2009). Winter canola is often planted in rotation 

with wheat, so weed control practices are applied to some extent ahead of the canola planting (e.g., 

Brown et al., 2008), which could limit the likelihood for presence of wild relatives in and around 

prospective winter canola fields. 
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Figure 19. Production Climate Zones for Spring, Biennial, and Winter Canola 

Map of climatic zones suitable for the production of different canola varieties (adapted from Mills, 1996) 

D2. Crop Relatives of Brassica napus 

The genus Brassica contains several important crop species that are used for a variety of purposes. 

Brassica napus and B. rapa are the most important for their use as oilseed crops (Ellstrand, 2003a). 

Brassica and the related genus Raphanus include vegetables for human consumption, e.g., swede (B. 

napus), turnip and Chinese cabbage (B. rapa), and cauliflower, cabbage, broccoli, and others (B. 

oleracea), Indian mustard (B. juncea), and radish (R. sativus). Three Brassica species and one species 

in the related genus Sinapis are ‘mustards’: B. carinata (Ethiopian mustard), B. juncea (Indian mustard), 

B. nigra (black mustard) and Sinapis alba (white mustard) (FitzJohn et al., 2007). Cultivation of B. 

carinata as an oilseed and vegetable crop is largely restricted to Ethiopia and India (Hemingway, 1995; 

Stewart, 2002). Some forms of B. napus, B. oleracea, B. rapa and R. sativus are also grown as fodder 

crops (FitzJohn et al., 2007). 

The three Brassica species forming the foundation of the Triangle of U (Nagaharu, 1935) are B. rapa, B. 

nigra, and B. oleracea. These three species are diploid, and their respective genomes have been 

distinguished from one another and are described as A, B, and C (Østergaard and King, 2008). They can 

be hybridized sexually with varying degrees of difficulty. Figure 20 describes the likelihood for B. napus

crossing successfully with other cultivated species. As B. napus and B. rapa may easily cross, other 

crosses with B. napus are difficult to nearly impossible based on genome characteristics (Myers, 2006; 

Nagaharu, 1935; OGTR, 2002). The resulting amphidiploid hybrids, B. juncea, B. napus, and B. carinata

contain AB, AC, and BC genomes, respectively. Research has shown that B. napus and B. rapa outcross 

readily with each other; these species can also outcross, albeit more rarely, with a wide range of wild and 

cultivated species including B. oleracea, B. nigra, and R. sativus (summarized in OGTR, 2002). Brassica 

napus and B. juncea share a common set of chromosomes, enhancing the likelihood of interspecific 

hybridization and gene flow (Myers, 2006).

 

Brassica napus and B. juncea can self-fertilize (Rakow and Woods, 1987), but B. rapa is selfincompatible (Warwick et al., 2003). The A genome is common to the three major oilseed Brassica

species, explaining the success of interspecific crossing, and the ability to transfer genes among these 

species (Figure 20). 

Canola 

Production Zone 

Spring Annual 

Biennial 

Winter Annual 
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Figure 20. Genome Relationships Among Cultivated Brassicaceae 

A modified “Triangle of U” showing genome relationships among cultivated Brassicaceae adapted from Nagaharu 

(1935) and Myers (2006). Genomes are represented by letters, and haploid genome chromosome numbers are 

described in parentheses. The three diploid species possess three basic genomes, A, B, and C. Lines represent 

ease with which species can be crossed. These genomes are combined in the three tetraploid species in pair-wise 

combinations, i.e. AB, AC, and BC. 

D3. Wild Relatives of Brassica napus 

Hybridization between crops and their wild relatives (either the same or different species) has occurred in 

agricultural history (Ellstrand and Schierenbeck, 2000; Raybould and Wilkinson, 2005). The majority of 

important crops have wild relatives (CAST, 2007; Ellstrand et al., 1999), and gene flow to wild species 

from crops occurs at higher frequency than might be expected (Ellstrand et al., 1999). However, the 

probability of gene flow from crops to related weed species is habitat-dependent and variable between 

populations (Cureton et al., 2006). 

The Brassicaceae family contains a number of major weeds, including those in the genera Sinapis, 

Capsella, Thlaspi, Erucastrium, Raphanus, and others (OECD, 1997). Concerns have been raised about 

the potential for the transfer of transgenes from the cultivated oilseed Brassica species to their weedy 

relatives in Europe and North America where Brassica crop species are widely grown. Some Brassica

crops and their wild relatives will hybridize only under artificial conditions in laboratories or highly 

contrived field conditions; others will hybridize at very low rates under natural conditions (Table 26; Barton 

and Dracup, 2000; Raybould, 1999). However, major barriers to introgression exist, including spatial and 

temporal isolation, sexual incompatibility, or lack of fertile progeny, thereby reducing the likelihood of 

gene flow. 

Table 26 describes crop and wild relative species that occur in the U.S. to some extent, and the likelihood 

for hybridizations with canola. In order to be conservative in the analysis of possible gene flow, species 

were considered for further examination if the species had geographical prevalence in canola production 

areas and at least one reported successful hybridization (hand pollination or in the field) with B. napus. If 
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information was not known about hybridization, the species was also examined further if present in 

primary canola-growing areas. The species that met these criteria are highlighted in Table 26 and 

discussed individually in section D5. 

Table 26. List of Species with Reports of Hand-Pollinated or Spontaneous Hybridization with 

B. napus

Species Namea Crop Weed 

Classificationb

Hand Pollination 

(successes:failures)c

Spontaneous and 

Natural 

Hybridizationc

Presence in 

Winter 

Canola 

Growing 

Areasd

Presence 

in Spring 

Canola 

Growing 

Areasd B.napus

as Male 

B.napus

as 

Female 

B.napus

as Male 

B.napus

as 

Female 

Brassica carinata  X 4:1 7:0  Brassica elongata   

Brassica fruticulosa  0:1 1:1  X

Brassica juncea   25:1 13:4  

Brassica nigra   2:2 4:2 X X  

Brassica oleracea  X 3:11 9:17  X  

Brassica rapa   55:8 84:0    

Brassica tournefortii  0:1 1:1  X

Camelina sativa  X 0:1 0:1  

Capsella bursapastoris  0:1 0:1  

Coincya monensis   

Conringia orientalis  0:1 0:1  

Diplotaxis erucoides X 1:1  

Diplotaxis muralis  3:0 1:1  

Diplotaxis siifolia X 0:3 0:1 X X

Diplotaxis tenuifolia  0:3 1:1  

Eruca vesicaria 

(E.sativa)  2:0  

Erucastrum gallicum  0:1 1:0 X  

Hirschfeldia incana  1:2 1:2    X

Moricandia arvensis X 0:2 0:2 X X

Myagrum perfoliatum X 0:1 0:1  X

Raphanus 

raphanistrum  0:4 3:2    

Raphanus sativus   1:5 1:2  

Rapistrum rugosum  1:0  

Rorippa islandica  1:0 X X

Sinapis alba   0:6 1:2 X  

Sinapis arvensis  1:10 5:8 X   

Sisymbrium irio  0:1 0:1  X

Sisymbrium orientale   

 = Yes, to some degree 

X = No 

a Species highlighted in purple have at least one report of successful hybridization with B. napus b

 Weed Science Society of America or USDA NRCS list of noxious weeds c

 Andersson and de Vicente, 2010; FitzJohn et al., 2007 d

 USDA PLANTS Database; Criteria to distinguish growing areas for winter and spring canola were 

obtained from Brown et al. (2008) and are summarized in the following section. 

Blank cells = no data available 
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D4. Gene Flow Assessment Considerations 

In general, it is difficult for a transgene to become established in a wild relative or recipient population. In 

order for successful introgression of a transgene in a recipient population, several conditions must exist 

and several steps must take place before this would occur (Devos et al., 2009). These are as follows: 

1. There must at least be a partial overlap in flowering periods between the transgenic canola and 

the relative. 

2. The relative must grow within the physical range of viable pollen dispersal of the transgenic 

canola, and viable pollen grains must reach the recipient stigma. 

3. Viable and fertile interspecific F1 hybrids must be produced, and the transgene successfully 

integrated into the recipient plant genome. 

4. The transgene must be transmitted through successive backcross generations. 

5. The transgene must be stabilized in the genome of the relative through backcrossing over 

multiple generations. 

6. The introgressed transgene must be either neutral or provide a fitness advantage such that it 

persists in the relative’s populations. 

In order to evaluate the likelihood of gene flow of 73496 canola to the species highlighted in Table 26, 

additional information related to canola flowering synchrony and flowering characteristics, pollen dispersal 

mechanisms and pollinator biology are discussed below in order to provide context for the gene flow 

assessment. 

Flowering synchrony and flower characteristics

Winter canola can grow in a broader geographic range than spring canola, thereby increasing the number 

of wild relative species with which geographic overlap is possible (Table 26). However, the flowering 

period may not overlap. Planting of winter canola occurs in the fall, and the plants overwinter in the 

rosette stage (Boyles et al., 2009; Boyles, 2010; Wysocki et al., 2005). Rosettes resume growth in the 

spring as temperatures increase, and (Boyles et al., 2009; Boyles, 2010; Wysocki et al., 2005) Depending 

on location, flowering for winter canola generally occurs in April to May, with harvest in June to July 

(Boyles et al., 2009; Boyles, 2010; Wysocki et al., 2005). As an example, Brassica tournefortii generally 

flowers from February to May in Arizona, in December to January in California, and January to March in 

Texas (summarized by Halvorson, 2003), thereby limiting the likelihood for flowering synchrony with 

winter canola. 

In self-compatible species (e.g. B. napus, B. juncea) (Rakow and Woods, 1987), pollination typically 

occurs prior to flower opening, thus limiting exposure of the recipient stigma to pollen from another flower 

or species. Finally, canola crops produce 5x1012 pollen grains per hectare (Chèvre et al., 1999). 

Therefore, due to a general lack of mobility of pollen, any pollen coming from outside the immediate field 

would be competing with this large volume of pollen (Salisbury, 2002), thereby decreasing the likelihood 

for successful pollination by pollen from outside the immediate field.

Pollen dispersal mechanisms

B. napus is primarily a self-pollinating crop, and the majority of large-scale outcrossing studies with fertile 

canola indicate outcrossing rates of <2% (Salisbury, 2002). However, worst-case estimates of 

outcrossing suggest levels between 12 to 47% (Becker et al., 1992; Kapteijns, 1993; Rakow and Woods, 

1987). The pollen, which is heavy and sticky, can also be transferred from plant to plant through physical 

contact with neighboring plants, via pollinator insects and, to a smaller extent, by wind (Cresswell, 1994; 

Cresswell et al., 2002; Williams, 1984). In general, windborne pollen may make little to no contribution to 

long distance pollination (OGTR, 2002). 

Approximately half of canola pollen travels less than three meters from the source; the vast majority of 

pollen travels less than 10 meters, with the amount of pollen decreasing as the distance from the pollen 

source increases (Scheffler et al., 1993; Thompson et al., 1999). Pollinating insects, in particular 
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honeybees and bumblebees, play a major role in B. napus pollination and are believed to be involved in 

the transfer of pollen over long distances (OGTR, 2002; Salisbury, 2002). 

Pollinator biology

Insects are the primary pollen vectors (Hayter and Cresswell, 2003), and insect foraging behavior is 

complex. The dynamics of bee-mediated pollen movement depend on a number of factors including 

spatial arrangement of plants, environmental conditions, plant density, availability of pollen, and the size 

and location of the receiving populations (Ellstrand et al., 1989; Klinger et al., 1992; Levin and Kerster, 

1969; Rieger et al., 2002). 

 

In situations with abundant flowers, such as in a cultivated field, individual honey bees generally collect 

nectar and pollen from flowers in the same or immediately adjacent plants (Cresswell, 1999; OGTR, 

2002; Pierre, 2001; Ramsay et al., 1999). Occasionally, bees may travel much further and some pollen 

transfer may occur over longer distances, for example a maximum flight distance of four kilometers has 

been reported in the literature (Ramsay et al., 1999; Thompson et al., 1999). However, the majority of bee 

flights are less than one meter, thereby limiting the likelihood of long distance pollen dispersal via insect 

pollinators (summarized by OGTR, 2002). 

Honeybees forage during daylight and are unlikely to carry viable pollen grains to impact fertilization 

beyond 12 hours (Kraai, 1962). Honeybees are sensitive to weather events and barometric pressure, 

and respond to these events by decreasing foraging distances (APHIS, 1998). The distance and success 

of pollen-mediated gene flow is dependent on its dispersal in space by either wind or insects, and on the 

length of time the pollen grain remains viable (OGTR, 2002). Canola pollen viability gradually decreases 

after four to five days in natural circumstances depending on environmental conditions, particularly 

temperature and humidity (Andersson and de Vicente, 2010; Rantio-Lehtimäki, 1995). 

Likelihood of hybridization

Reproductive compatibility among Brassica crops is complex. Because experimental hybridization 

studies are designed to optimize the likelihood of successful hybridization, they create bias toward 

positive reports of hybridization between species that may be unlikely to cross in natural conditions 

(FitzJohn et al., 2007). The ease with which a crop and its wild relatives can hybridize through manual 

cross-pollination reveals little about the potential influence of pre-pollination and other ecological barriers 

(Arnold, 1997; FitzJohn et al., 2007; Grant, 1994). 

Many studies have been conducted to gather experimental data to infer the likelihood of hybridization 

using a variety of techniques: experimental crosses (manual hand pollination), spontaneous crosses 

(non-assisted crosses under field conditions), and in vitro methods (e.g., embryo rescue). Crosscompatibility varies with the particular genotype used and with the polarity of the cross (i.e., dependent 

upon which species was the maternal parent; Arnold, 1997). Most combinations are unsuccessful and 

where crosses are successful, rates of hybrid production are typically very low (Table 26; FitzJohn et al., 

2007; Warwick and Black, 1993). 

Hybridization of B. napus with cultivated and wild Brassica species such as B. carinata, B.elongata, 

B.fruticulosa, B. incana, B.nigra, B. tournefortii, have been attempted and in most cases the production of 

hybrids requires considerable human intervention and is only successful if artificial hybridization 

techniques are used (e.g., embryo rescue) (Andersson and de Vicente, 2010). All resulting F1 hybrids 

are sterile (Andersson and de Vicente, 2010). Brassica carinata, B.nigra, and B. tournefortii are able to 

form hybrids with B. napus after hand pollination however, they have significant barriers to introgression 

such as pollen dehiscense prior to the flower opening, sexual incompatibility, reduced hybrid fertility, or 

sterility, making gene exchange with B. napus extremely unlikely (Andersson and de Vicente, 2010). 

Even among the closely related Brassica species (Nagaharu, 1935), hybridization with B. napus is highly 

unlikely. For example, although B. napus and B. rapa share the A genome, Scott and Wilkinson (1998) 

reported only 7% of B. rapa populations had any hybrids, indicating that 93% of B. rapa populations had 

no hybrids, and hybridization rates were low (0.4-1.5%) in field situations; further, less than 2% of hybrid 
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seedlings survived (Scott and Wilkinson, 1998). Other published hybridization rates are equally low, e.g. 

<3% for B. juncea (Bing et al., 1991; Jørgensen et al., 1996), <0.003% for R. raphanistrum (Chèvre et al., 

1999; Chèvre et al., 2000), <0.000034% for S. arvensis (Lefol et al., 1996). In many cases of attempted 

hybridizations with B. napus as male donor, no hybrids were produced (Salisbury, 2002). 

D5. Assessment of Gene Flow Potential from 73496 Canola to Relevant Species 

Gene flow assessments were conducted for those species highlighted in Table 26 on the basis of 

overlapping geographic ranges with canola production and reports of at least one successful hybridization 

(by hand pollination or in the field) with B. napus. If information was not known about hybridization, the 

species was also examined further if present in primary canola growing-areas. 

Crop Species: 

Brassica carinata (Ethiopian mustard) 

Although some reports of successful hand pollinations are reported for B. napus and B. carinata, no 

hybridizations in the field have been reported (Andersson and de Vicente 2010), and no distribution 

information is available for B. carinata in the U.S. (USDA Plants Database, 2010). Several barriers to 

introgression exist, e.g. reduced hybrid fertility or sterility (Andersson and de Vicente, 2010). 

Brassica juncea (Indian or brown mustard) 

B. juncea can occur throughout the U.S. (USDA Plants Database, 2010) and is both a crop and a weed. 

B. juncea (AABB genome) has a common set of chromosomes with B. napus (AACC genome), which 

enhances the likelihood of interspecific hybridization and gene flow (Andersson and de Vicente, 2010). 

Spontaneous hybridization has been reported at <3% for B. juncea (Bing et al., 1991; Jørgensen et al., 

1996), although hybridization is less successful when B. napus is the female parent (Jørgensen et al., 

1998). In general, F1 hybrids are rarely more fit than highly selected and well adapted counterparts, and 

low initial frequencies, reduced fitness and viability, and competitive disadvantage with respect to parents 

can lead to hybrid extinction (Wolf et al., 2001). The pollen and seed fertility of the F1 hybrids is typically 

less than 30% (Bajaj et al., 1986; Choudhary and Joshi, 1999; Frello et al., 1995; GhoshDastidar and 

Varma, 1999; Prakash and Chopra, 1988; Roy, 1980; Sacristán and Gerdemann, 1986), however 

spontaneous backcrossing progeny with improved fertility have been documented (Bing et al., 1991; Bing 

et al., 1996; Jørgensen, 1999). Backcrossing and subsequent gene introgression from B. napus to B. 

juncea could be expected, although infrequently (Andersson and de Vicente, 2010). 

Brassica nigra (Black mustard)

B. nigra can grow throughout regions of the U.S. (USDA Plants Database, 2010) and is classified as both 

crop and weed. Hybrids between Brassica nigra and B. napus have been reported to occur at a low 

frequency when produced under controlled conditions (Bing et al., 1991; Heyn, 1977). However, under 

field conditions no hybrids resulted from co-cultivation of B. nigra and B. napus (Bing et al., 1996). 

Brassica oleracea (e.g. cauliflower, cabbage, broccoli)

B. oleracea has a more limited distribution than other Brassica spp. (i.e. CA, CT, DC, GA, IA, IL, KY, LA, 

MA, MO, ME, MI, MD, MT, NM, NY, OH, OR, PA, RI, TX, VA, WA) (USDA Plants Database, 2010). 

Hybridization of B. napus with B. oleracea can be achieved by hand pollination (Eastham and Sweet, 

2002; Scheffler and Dale, 1994). Although B. oleracea and B. napus share a common set of 

chromosomes (Figure 19), the frequency of successful crosses is very low, and embryos often abort at 

early stages of development (Chiang et al., 1977; Honma and Summers, 1976; Mattsson, 1988). Thus 

far, no viable hybrid seeds have been obtained from crosses between B. napus and B. oleracea without 

the assistance of embryo rescue or ovule culture (Ayotte et al., 1987; Myers, 2006; Quazi, 1988; 

Takeshita et al., 1980). Spontaneous hybrids have been reported in wild B. oleracea populations in the 
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UK (Ford et al., 2006). No information is available thus far on the fertility of the natural hybrids reported 

(Andersson and de Vicente, 2010). 

Brassica rapa (Turnip, Chinese cabbage)

B. rapa is a closely related species of canola (amphidiploid), which has a similar life history to canola, but 

with a shorter growing season. B. napus (AACC) and B. rapa (AA) have a common set of chromosomes, 

making interspecific outcrossing more common (e.g. Bing et al., 1991). Gene flow measurements by Scott 

and Wilkinson (1998) reported only 7% of B. rapa populations had any hybrids, indicating that 93% of B. 

rapa populations had no hybrids, and hybridization rates were low (0.4-1.5%) in field situations. 

Importantly, less than 2% of hybrid seedlings survived (Scott and Wilkinson, 1998). 

Hybridization frequencies are higher when B. rapa occurs as a weed within canola crops, but varies 

significantly with experimental design. Where natural interspecific hybrids occur, hybrids have reduced 

fertility and low seed set (average 2-5 per pod) compared with the parents (Jørgensen and Andersen, 

1994). Reduced dormancy of B. rapa x canola hybrids relative to the persistent wild B. rapa (Jørgensen et 

al., 1999), coupled with the reduced fertility of the inter-specific hybrid (Jørgensen et al., 1999) makes it 

very unlikely that populations of these hybrids will persist. 

Raphanus sativus (radish) 

R. sativus can be found throughout the U.S. (USDA Plants Database, 2010). Spontaneous hybridization 

between a male sterile B.napus line and a cultivated variety of R. sativus line has been reported 

(Ammitzboll and Jørgensen, 2006). However, all F1 offspring were found to be hybrids with low pollen 

fertility (0%-15%) (Ammitzboll and Jørgensen, 2006). Although hybridization can be achieved by hand 

pollination or through sophisticated methods, such as ovule culture and embryo rescue (Gupta, 1997; 

Huang et al., 2002; Metz et al., 1995; Rhee et al., 1997; Sundberg and Glimelius, 1991) all artificially 

produced F1 hybrids have shown to be male sterile (Andersson and de Vicente, 2010). 

Sinapis alba (See Other Highlighted Species section below). 

Weed Species: 

Brassica fruticulosa

This species only occurs in California (USDA Plants Database, 2010), therefore risk is limited to canola 

production in California. Additionally, only one report of successful hand pollination has occurred, and no 

reports of hybridization in the field are described (Andersson and de Vicente, 2010). 

Brassica tournefortii

B. tournefortii grows in the southwestern U.S. (e.g. NV, NM, AZ, TX, CA) (USDA Plants Database, 2010), 

therefore geographic overlap is limited to winter canola production, however as previously discussed, 

asynchrony of flowering is probable in this case (Boyles et al., 2009; Boyles, 2010; Halvorson, 2003; 

Wysocki et al., 2005). 

Hybridization of B. napus has been attempted with B. tournefortii. In most cases, the production of F1 

hybrids requires considerable human intervention and is only successful if artificial hybridization 

techniques are used. B. tournefortii is able to form hybrids with B.napus after hand pollination. However, 

they have significant barriers to introgression such as pollen dehiscence prior to the flower opening, 

sexual incompatibility, reduced hybrid fertility, or sterility making gene exchange with B.napus extremely 

unlikely (Andersson and de Vicente, 2010) 
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Diplotaxis muralis

D. muralis occurs throughout regions of the U.S. (USDA Plants Database, 2010). Ringdahl et al. (1987) 

examined the crosssability of D. muralis and B. napus using conventional crossing techniques, using B. 

napus as the male parent. They were able to produce viable F1 hybrids after hundreds of hand 

pollinations conducted (Ringdahl et al., 1987). On the other hand, Fan et al. 1985 investigated male 

sterility in backcross populations from hybrids between D. muralis and B.napus, using D. muralis as the 

female parent. Male sterility of the F1 populations was caused by an extra chromosome derived from 

D.muralis (Fan et al., 1985). 

Erucastrum gallicum

E. gallicum can be found throughout regions of the U.S. (USDA Plants Database, 2010), however studies 

conducted by Warwick et al. (2003) indicated that gene flow from commercial fields of B. napus to E. 

gallicum has very low probability of occurrence. 

Hirschfeldia incana

H. incana can be found in CA, HI, NV, OR (USDA Plants Database, 2010). Spontaneous hybridization 

between male-sterile B. napus and H. incana in field has been reported in both directions (Chèvre et al., 

1996; Eber et al., 1994; Lefol et al., 1995). The resulting triploid F1 hybrids are often vigorous and at least 

as competitive as their wild parent (Eber et al., 1994; Lefol et al., 1996). However, they are usually male 

sterile and produce very little seed (1 seed per plant) under controlled conditions (Lefol et al., 1996). 

Hybridization between these two species is easy and recurrent, but introgression rarely occurs (Darmency 

and Fleury, 2000). 

Raphanus raphanistrum

R. raphanistrum occurs throughout regions of the U.S. (USDA Plants Database, 2010), and there is a 

possibility for successful hybridization; however, the frequency of such hybridization is very low (e.g. 

<0.003%; Ammitzboll and Jørgensen, 2006; Baranger et al., 1995; Chèvre et al., 1996; Chèvre et al., 

1999; Chèvre et al., 2000; Darmency et al., 1995; Eber et al., 1994; Rieger et al., 2001;), and hybrids 

show very low fertility (Andersson and de Vicente, 2010). 

Spontaneous hybridization between B. napus and R. raphanistrum may occur in the wild. Low 

frequencies of F1 hybrids have been reported and they are allotriploids, which show very low fertility 

(Baranger et al., 1995; Chèvre et al., 1996; Chèvre et al., 1998; Darmency et al, 1998; Kerlan et al., 1992; 

Pinder et al., 1999; Thalmann et al., 2001; Warwick, 2009). F1 hybrids also commonly show decreased 

fitness in terms of reduced seedling emergence, a significant emergence delay, and lower survival rate 

than both parents. (Guéritaine et al., 2003). 

Rorippa islandica

Although intergeneric hybridization between R. islandica and B. napus is possible (Bijral and Sharma, 

1995), the distribution of this species in the U.S. is restricted to Washington D.C. (USDA Plants Database, 

2010) therefore the potential for gene flow and hybridization is negligible. 

Sinapis arvensis

S. arvensis occurs throughout the U.S. (USDA Plants Database, 2010) and is a weed frequently found in 

canola fields in weed surveys (Zollinger et al., 2003). However, greenhouse studies conducted by Moyes 

et al. (2002) have confirmed the low probability of hybridization between B. napus and S. arvensis. Low 

frequencies (<0.00003%) of hybrids were obtained when S. arvensis was the receptor (female) (Lefol et 

al., 1996). Although a low level of interspecific crosses could occur (Leckie et al., 1993), the likelihood of 

gene flow between B. napus and S. arvensis is extremely remote, and there is general agreement that no 

gene introgression will occur (Bing et al., 1991; Downey, 1999; Eber et al., 1994). Hybrids obtained 
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between S.arvensis and B. napus (Moyes et al., 2002) had greatly reduced fertility or were completely 

sterile (Andersson and de Vicente, 2010). 

Other Highlighted Species 

Of the remaining highlighted species (Table 26), C. monesis occurs in CA and the eastern U.S. in states 

with very limited canola production (USDA Plants Database, 2010). B. elongata is present in CO, NV, and 

OR, and S. orientale occurs in the southwestern U.S. (USDA Plants Database, 2010). Crosses between 

B. napus and a number of species belonging to other genera have been attempted using artificial 

hybridization methods, but the crosses either failed or resulted in partially or completely sterile F1 

progeny. Among those species are: Diplotaxis erucoides, Diplotaxis tenuifolia, Eruca vesicaria (E. sativa), 

Erucastrum gallicum, Rapistrum rugosum, Sinapis alba, Sisymbrium orientale (Andersson and de 

Vicente, 2010). 

Related Weeds Frequently Found in Agricultural Fields

In addition to examination of geographic range and likelihood of hybridization, it is important that 

abundance of weed species in agricultural fields is considered in the risk assessment. Weed abundance 

in agricultural fields is partially determined by crop production practices, and weed species prominence 

can be significantly impacted by cropping systems and cultivation practices (CFIA, 1994). In North 

Dakota weed surveys, some Brassicaceae weed species that are frequently found include Thlaspi 

arvense, Descurainia sophia, and Descurainia pinnata (Zollinger et al., 2003). The probability of 

hybridization and gene flow to these species is unknown. 

D6. Conclusions on the Potential Plant Pest Risk of 73496 Canola 

73496 canola does not pose a plant pest risk. The GAT4621 protein and its source organism, B. 

licheniformis, are familiar and have a history of safe use. The transgenic DNA in 73496 canola is as safe 

for consumption as any other DNA. B. napus is not considered a noxious or invasive weed in the U.S. 

Although it does demonstrate some weedy characteristics, canola has undergone many years of selective 

breeding and domestication and is a poor competitor with other species. In addition, the agronomic 

characteristics and germination data for 73496 canola demonstrated no change that would allow for 

development of weedy characteristics different from other canola varieties. The gat4621 gene does not 

provide a selective advantage in unmanaged ecosystems, rather only in settings where glyphosate is 

being applied for weed control. Other commercially available herbicides, such as glufosinate or 

imidazolinones, as well as mechanical means can be used in crop settings for volunteer control. 

Gene flow from 73496 canola was evaluated thoroughly with respect to plant pest risk. Although crops 

and certain wild/weedy relatives have exchanged genes for centuries, the concern with genetically 

modified crops is that the acquisition of transgenes may increase the fitness of recipient plants (Ellstrand 

et al., 1999; Ellstrand, 2003b) and the potential weediness or invasiveness in the crop itself or in its wild 

or weedy relatives as a result of transgene movement (Warwick et al., 2009). Successful hybridization of 

B. napus and a wild/weedy relative is highly unlikely. However, should such unlikely events of successful 

hybridization and stable introgression of the gat4621 gene from 73496 canola into wild/weedy relative 

populations occur, the herbicide-tolerance trait would only provide selective advantage in situations in 

which the weedy hybrid was in contact with the herbicide (i.e., in an agricultural field). 

The introduced gat4621 gene in 73496 canola did not change the ability of the plant to interbreed with 

other plant species. Furthermore, the evaluation of agronomic and phenotypic properties of 73496 

canola, including those characteristics associated with reproductive biology, indicated no unintended 

changes likely to affect the potential for gene flow from 73496 canola to sexually compatible species. 

The consequences of gene flow and introgression of the glyphosate-tolerant trait from 73496 canola to 

the same or sexually compatible species is anticipated to be the same as for existing commercial 

glyphosate-tolerant canola varieties. If glyphosate-tolerant individuals arise through interspecific or 

intergeneric hybridization, the tolerance will not confer any competitive advantage to these plants unless 
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selected by glyphosate herbicide. This would only occur in managed ecosystems where glyphosatecontaining herbicides are applied for broad-spectrum weed control, or in plant varieties developed to 

exhibit glyphosate tolerance and in which glyphosate is used to control weeds. As with glyphosatetolerant canola volunteers, these individuals, should they arise, would be controlled using other available 

chemical or mechanical means. 

Although gene flow from 73496 canola to relatives is possible, it will not result in increased weediness or 

invasiveness of these relatives based on the agronomic and ecological assessments (Section VII; 

Appendix 5). Large-scale cultivation of herbicide-tolerant canola has occurred for nearly 15 years in 

Canada and the United States. To date, there are no reports of problems with interspecific crosses and 

introgression of herbicide-tolerant genes into cultivated or wild relatives of canola (Andersson and de 

Vicente, 2010). 
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Appendix 1. Materials and Methods for Genetic Characterization of 73496 Canola 

1.1. Southern Blot Characterization of 73496 Canola 

Southern blot analysis was conducted to characterize the DNA insertions in 73496 canola. Individual 

plants of the T2, T3, T3F2, T3F3, and F1*2 generation were analyzed by Southern blot to determine the 

number of each of the genetic elements of the expression cassette that were inserted and to verify that 

the integrity of the PHP28181A fragment was maintained upon integration. The integration patterns of 

the insertion in 73496 canola was investigated with Nco I and Ssp I restriction enzymes. Southern blot 

analysis was conducted on individual plants of the five generations to confirm stability of the insertion 

across generations and to verify the absence of backbone sequences from plasmid PHP28181. 

1.1.1. Test Material 

Seeds from the T2, T3, T3F2, T3F3, and F1*2 generations of 73496 canola were planted and leaf tissue 

harvested from individual plants was used for genomic DNA extraction. 

1.1.2. Control Material 

Seeds from the unmodified canola varieties 1822B, 1822R, and 6395B were planted and leaf tissue 

harvested from individual plants was used for genomic DNA extraction. 

1.1.3. Reference Material 

Plasmid DNA from PHP28181 was prepared from E. coli (Invitrogen, Carlsbad, CA) and was used as a 

positive control for Southern blot analysis to verify probe hybridization and to verify sizes of fragments 

internal to the plasmid. The plasmid stock was a copy of the plasmid used for transformation to produce 

73496 canola and was digested with restriction enzymes to confirm the plasmid map. The probes used in 

this study were derived from plasmid PHP28181 or from a plasmid containing equivalent genetic 

elements. 

DNA molecular weight markers for gel electrophoresis and Southern blot analysis were used to determine 

approximate molecular weights. For Southern analysis, DNA Molecular Weight Marker VII, digoxigenin 

(DIG) labeled (Roche, Indianapolis, IN), was used as a size standard for hybridizing fragments. ΦX174 

RF DNA/Hae III Fragments (Invitrogen, Carlsbad, CA) was used as a molecular weight standard to 

determine sufficient migration and separation of the fragments on the gel. 

1.1.4. Genomic DNA Extraction 

Genomic DNA was extracted from leaf tissue harvested from individual plants as described above. The 

tissue was pulverized in tubes containing grinding beads using a Geno/GrinderTM (SPEX CertiPrep, Inc., 

Metuchen, NJ) instrument and the genomic DNA was isolated using a urea-based procedure 

(modification from Chen and Dellaporta, 1994). Approximately 1 gram of ground tissue per sample was 

extracted with 5 ml Urea Extraction Buffer (7 M urea, 0.34 M NaCl, 0.05 M Tris-HCl, pH 8.0, 0.02 M 

EDTA, 1% N-lauroylsarcosine) for 15-18 minutes at 37C, followed by two extractions with 

phenol/chloroform/isoamyl alcohol (25:24:1) and one extraction with water saturated chloroform. The 

DNA was precipitated from the aqueous phase by the addition of 1/10 volume of 3 M NaOAc (pH 5.2) and 

1 volume of isopropyl alcohol, followed by centrifugation to pellet the DNA. After washing the pellet twice 

with 70% ethanol, the DNA was dissolved in 0.5 ml distilled water and treated with 10 g ribonuclease A 

for 15 minutes at 37C. The sample was then washed with 70% ethanol. After drying, the DNA was redissolved with 0.5 ml distilled water and stored at 4C.
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1.1.5. Quantitation of Genomic DNA 

Following extraction, the DNA samples were quantified on a spectrofluorometer using PicoGreen® 

reagent (Molecular Probes, Inc., Eugene, OR) following a standard procedure. The DNA was also 

visualized on an agarose gel to confirm quantitation values from the PicoGreen® analysis and to 

determine DNA quality. 

1.1.6. Identification of the 73496 Canola Plants Used for Southern Blot Analysis 

Phenotypic analysis of 73496 canola plants and control plants was carried out by the use of lateral flow 

devices able to detect the GAT4621 protein to confirm the absence or presence of the GAT4621 protein 

in material used for Southern blot analysis. 

Leaf extract were prepared by grinding leaf punches to homogeneity in 400 μl of SEB6 extraction buffer 

(Agdia, Inc., Elkhart, IN). Lateral flow devices (Agdia) were placed in the homogenate and allowed to 

develop. After incubation, the results were read from the lateral flow devices. A single stripe indicated a 

negative result and a double stripe indicated the sample was positive for the GAT4621 protein. 

Genotypic analysis of the 73496 canola and control canola plants was carried out by real-time 

polymerase chain reaction (PCR) using assays specific for the DNA insertion. A leaf sample was taken 

from each test and control plants for event-specific PCR analysis. DNA was extracted from each leaf 

sample using the Extract-N-AmpTM Plant PCR kit using the described procedure (Sigma-Aldrich, St. 

Louis, MO). 

Real-time PCR was performed on each DNA sample utilizing an ABI PRISM 7500 Fast Real-Time PCR 

System (Applied Biosystems, Inc., Foster City, CA). TaqMan®

 probe (Applied Biosystems, Inc.) and 

primer sets (Integrated DNA Technologies, Coralville, IA) were designed to detect target sequences from 

the insertion in 73496 canola. In addition, a second TaqMan®

 probe and primer set for a reference canola 

endogenous gene was used to confirm the presence of amplifiable DNA in each reaction. The assay 

analysis consisted of real-time PCR determination of qualitative positive/negative calls. The extracted 

DNA was assayed using TaqMan®

 Universal PCR Master Mix, No AmpErase® UNG (Applied Biosystems, 

Inc.). Initial incubation was at 95C for 10 minutes followed by 40 cycles as follows: 95C for 15 seconds, 

60C for one minute. 

Positive or negative determination for each plant was based on comparison of the threshold cycle (CT) of 

the insertion target PCR to that of the canola endogenous reference target. If the event-specific and 

endogenous PCR targets amplified above CT, then the plant was scored as positive for the DP-Ø73496-4 

event. If the endogenous target amplified and the event target did not, then the plant was scored as 

negative. For all assays, if neither target amplified for a particular sample, it was determined to be a poor 

quality sample or failed run and the assay was repeated. 

A subset of 73496 canola plants that were identified as containing the inserted DNA and expressing the 

GAT4621 protein from the five generations described previously were selected for Southern blot analysis. 

1.1.7. Digestion of DNA for Southern Blot Analyses 

Genomic DNA samples extracted from selected 73496 canola and control canola plants were digested 

with restriction enzymes following a standard procedure. Approximately 4 g of genomic DNA was 

digested using 20-50 units of enzyme according to manufacturer’s recommendations. The digestions 

were carried out at 37C for approximately three hours, followed by ethanol precipitation with 1/10 volume 

of 3 M NaOAc (pH 5.2) and 2 volumes of 100% ethanol. After incubation at ≤-5C and centrifugation, the 

DNA was allowed to dry and then re-dissolved in TE buffer (10mM Tris, 1 mM EDTA, pH 7.5). The 

reference plasmid, PHP28181, was spiked into a control plant DNA sample in an amount equivalent to 

approximately one or three gene copies per canola genome and digested with the same enzyme to serve 
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as a positive control for probe hybridization and to verify sizes of fragments internal to the plasmid on the 

Southern blot. 

1.1.8. Electrophoretic Separation and Southern Transfer 

Following restriction enzyme digestion, the resultant DNA fragments were electrophoretically separated 

by size through an agarose gel. A molecular weight standard [ΦX174 RF DNA/HaeIII Fragments 

(Invitrogen)] was used to determine sufficient migration and separation of the fragments on the gel. DIG 

labeled DNA Molecular Weight Marker VII (Roche), which is visible after DIG detection as described 

below, was used to determine hybridizing fragment size on the Southern blots. 

Agarose gels containing the separated DNA fragments were depurinated, denatured, and neutralized in 

situ, and transferred to a nylon membrane in 20x SSC buffer (3M NaCl, 0.3 M sodium citrate) using the 

method as described for the TURBOBLOTTERTM Rapid Downward Transfer System (Whatman, Inc., 

Piscataway, NJ). The DNA was then bound to the membrane by UV crosslinking (Stratalinker, 

Stratagene, La Jolla, CA). 

1.1.9. DNA Probe Labeling for Southern Blot Hybridization 

Probes for the UBQ10 promoter, gat4621, and pinII terminator were used to detect genes and elements 

within the insertion. Probes covering the backbone region of plasmid PHP28181 were used to verify 

absence of plasmid backbone DNA in 73496 canola. DNA fragments of the probe elements were 

generated by PCR from plasmid PHP28181 or a plasmid with equivalent elements using specific primers. 

PCR fragments were electrophoretically separated on an agarose gel, excised and purified using a gel 

purification kit (Qiagen, Valencia, CA). DNA probes were generated from these fragments by PCR that 

incorporated a DIG labeled nucleotide, [DIG-11]-dUTP, into the fragment. PCR labeling of isolated 

fragments was carried out according to the procedures supplied in the PCR DIG Probe Synthesis Kit 

(Roche). 

1.1.10. Probe Hybridization and Visualization 

Labeled probes were hybridized to the target DNA on the nylon membranes for detection of the specific 

fragments using the procedures essentially as described for DIG Easy Hyb solution (Roche). After 

stringent washes, the hybridized DIG-labeled probes and DIG-labeled DNA standards were visualized 

using CDP-Star Chemiluminescent Nucleic Acid Detection System with DIG Wash and Block Buffer Set 

(Roche). Blots were exposed to X-ray film for one or more time points to detect hybridizing fragments and 

to visualize molecular weight standards bound to the nylon membrane. Images were digitally captured by 

detection with the Luminescent Image Analyzer LAS-3000 (Fujifilm Medical Systems, Stamford, CT). 

Digital images were compared to original X-ray film exposures as verification for use in this submission. 

The sizes of detected bands were documented for each digest and each probe. 

1.1.11. Stripping of Probes and Subsequent Hybridizations 

Following hybridization and detection, membranes were stripped of DIG-labeled probe to prepare the blot 

for subsequent re-hybridization to additional probes. Membranes were rinsed briefly in distilled, deionized water and then stripped in a solution of 0.2 M NaOH and 0.1% SDS at 37°C with constant 

shaking. The membranes were then rinsed in 2x SSC and used directly for subsequent hybridizations. 

The alkali-based stripping procedure effectively removes probes labeled with the alkali-labile DIG.
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1.2. Mendelian Inheritance and Genetic Stability of the Trait in 73496 Canola

Inheritance of the inserted DNA and the herbicide-tolerance phenotype in 73496 canola was evaluated to 

ensure stability of the trait during the plant breeding process and to confirm the trait was at a single 

genetic locus. A phenotypic and genotypic analysis of individual plants of segregating and nonsegregating generations was conducted. The presence of the DP-Ø73496-4 event insertion was 

determined by event- and gat4621 gene-specific endpoint PCR analysis performed on leaf punches from 

seedlings of each generation. Plant phenotypic analysis was conducted by treating the plants with 

herbicide and determining the level of herbicide injury to the plants. 

1.2.1. Test Material 

One-hundred plants from segregating generations of 73496 canola (T3F2, BC1F1*1,*3, BC2F1*1,*3, and 

BC3F1*1,*3) and one non-segregating generation (F1*1,*3) were evaluated. The breeding history of these 

five generations is shown in the breeding diagram in Section III-A, Figure 2. Those populations with the 

superscript “*1,*3” designation were pooled populations from two different genetic backgrounds (Section 

III-A, Figure 2). The plants from the T3F2 generation were expected to segregate 3:1 (positive:negative 

for the presence of the inserted DNA and the herbicide-tolerance phenotype), and the plants from the 

BC1F1, BC2F1, and BC3F1 generations were expected to segregate 1:1 (positive:negative). Leaf tissue 

samples were collected for conducting the genotypic analysis described below. 

1.2.2. Genotypic Analysis 

Leaf punch samples collected from 73496 canola plants were analyzed using qualitative endpoint PCR 

analysis to confirm the presence or absence of the DP-Ø73496-4 event insertion and the gat4621 gene. 

The PCR results for one 73496 canola plant (generation T3F2) were inconclusive and the plant had 

already been discarded so no plant material was available for retesting; therefore, the PCR results for this 

plant was not included in subsequent statistical analysis. 

1.2.3. Phenotypic Analysis 

After sample collection, all plants were treated with a mixture containing the herbicide Touchdown 

HiTecha

 (Syngenta Crop Protection, Inc., Greensboro, NC), non-ionic surfactant, and ammonium sulfate. 

Glyphosate constitutes 52.3% (by weight) of Touchdown HiTech in the form of a monopotassium salt, 

equivalent to 5 pounds glyphosate acid equivalents per gallon (0.6 kilograms glyphosate acid equivalents 

per liter). The spray mixture was applied at an approximate rate of 16 fluid ounces per acre (1.2 liters per 

hectare) of Touchdown HiTech, 3.5 pounds per acre (3.9 kilograms per hectare) of ammonium sulfate, 

and 0.25% (by volume) of non-ionic surfactant per volume of spray solution. Spray volume was 

approximately 23 gallons per acre (215 liters per hectare). 

Each plant was visually evaluated 9 days after herbicide application for the presence of herbicide injury, 

and was identified as presenting an herbicide-tolerant phenotype (plant exhibited no herbicidal injury) or 

an herbicide-susceptible phenotype (plant exhibited severe herbicide injury). After herbicide injury 

evaluation, all plants were discarded. 

1.2.4. Statistical Analysis of Trait Inheritance Data 

Statistical analyses were conducted using SAS software, Version 9.2 (SAS Institute, Inc., Cary, NC). A 

Pearson’s chi-square (χ2

 test at 95% confidence) was conducted for each segregating generation of 

73496 canola to determine the goodness-of-fit of the observed segregation ratio to the expected 

segregation ratio. This value was not calculated for the non-segregating populations. The Pearson test 

statistic was conducted separately for both the genotypic (i.e., PCR) results and phenotypic (i.e., 

herbicide tolerance) results, and was computed using the following equation: 

 

a Registered trademark of a Syngenta Group Company
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73496 Canola 

2 2

2 ( )( ) pos pos neg neg

pos neg

OE OE

E E     

Where Opos denotes the total observed positive frequencies, Epos denotes the total expected positive 

frequencies, Oneg denotes the total observed negative frequencies, Eneg denotes the total expected 

negative frequencies, and χ2 follows a chi-square distribution with one degree of freedom (df). 

 

A chi-square value >3.84 (χ2

(df=1)) has a P-value <0.05 and indicates a significant difference between 

the observed and expected frequencies. A chi-square value ≤3.84 (χ2(df=1)) has a P-value ≥0.05 and 

indicates that the observed data can be interpreted as consistent with the expected ratio. 
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Appendix 2. Description of Statistical Analyses Performed for Agronomic and Nutrient 

Composition Data 

2.1. Statistical Analysis of Germination Data 

Statistical analyses were conducted to evaluate and compare germination rates of 73496 canola and a 

near-isoline control canola using version 9.1 of SASb

 software (SAS Institute, Inc., 100 SAS Campus 

Drive, Cary, NC 27513, USA). Statistical analyses were conducted separately for each of three 

germination tests (warm, cold, and diurnal). 

2.1.1. Mixed Model for Statistical Analysis 

For a given germination test, data were analyzed using a Generalized Linear Mixed Model (GLMM), 

assuming Binomial distribution with the “logit” link function: 

GLMM model on the linear predictor scale: ij = i

 + rj(i) Model 1

 rj(i) iid ~ N(0,

2

R)

where μi denotes the mean response for the ith entry (fixed effect) and rj(i) denotes the effect of the jth replication within 

the ith entry (random effect nested within fixed effect), and iid denotes independently and identically distributed. 

Binomial Distribution: yij|rj(i) ~ Binomial (nij,ij) 

where yij denotes the number of germinated seed in the jth replication of the ith entry, j = 1, 2, ..., 8 (each replication 

contained a total of 46~72 seed), nij denotes the total number of seed in the jth replication of the ith entry, and ij

denotes the probability of a seed being germinated in the jth replication of the ith entry. 

“Logit” link function: logit( ) log 1

ij

ij ij

ij



 



          

which transformed the parameter ij into a linear predictor, denoted as nij. Logit link is default for Binomial data. 

2.1.2. Least Squares Mean (LS-Mean) Calculations 

For a given germination test, the mean (i.e. least squares mean, also referred to as LS-Mean) value and 

standard error was estimated from Model 1 for each of 73496 canola and the control canola. 

2.1.3. Statistical Comparisons and Interpretations 

To test for differences in LS-Mean values between 73496 canola and the control canola, a p-value was 

calculated for each germination test. For a given comparison, a p-value of < 0.05 was considered to 

indicate a significant difference. 

2.2. Statistical Analyses of Agronomic Characteristic, Yield, and Nutrient Composition Data 

Statistical analyses were conducted to evaluate and compare agronomic characteristics and yield of 

73496 canola and near-isoline control canola across seven sites during the 2008 growing season 

(referred to as Experiment A), and across three sites during the 2009 growing season (referred to as 

Experiment B). Statistical analyses were also conducted to evaluate and compare nutrient composition of 

seed derived from 73496 canola and near-isoline control canola across six sites during the 2009 growing 

season. 

 

b

 Registered trademark of the SAS Institute, Inc. 
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Analysis was conducted separately for each study using SAS software (version 9.1 of SAS was used for 

analysis of Experiment A, and version 9.2 was used for Experiement B and the nutrient composition 

data). Days to maturity data were not collected at one site and early growth data were not collected at 

four sites for Experiment A. Yield data collected at one site were not included in the respective analysis 

for Experiment B. 

2.2.1. Mixed Models for Across Sites Analysis 

Default Model

For a given agronomic characteristic or composition analyte, data were analyzed using the following 

linear mixed model: 

yijk = μi

 + lj + rk(j) + (μl)ij + εijk Model 2 

lj

 ~ iid N(0, σ2

Site), rk(j) ~ iid N(0,σ2Rep), (μl)ij ~ iid N(0, σ2Ent×Site), and εijk ~ iid N(0, σ2Error)

where μi denotes the mean of the ith entry (fixed effect), lj denotes the effect of the jth site (random effect), rk(j) denotes 

the effect of the kth block within the jth site (random effect), (μl)ij denotes the interaction between the entries and sites 

(random effect) and εijk denotes the effect of the plot assigned the ith entry in the kth block of the jth site (random effect 

or residual). Notation ~iid N(0, σ2

a) indicates random variables that are identically independently distributed (iid) as 

normal with zero mean and variance σ2

a. 

 

Residual maximum likelihood (REML) estimation procedure was used to generate estimates of variance 

components, and entry means (i.e. LS-Means) across sites were then estimated with standard errors. 

Model Allowing Negative Covariance

When the REML estimate for the entry by site variance component σ2

Entry×Site from Model 2 was zero, the 

random part of the mixed model was changed to a compound symmetry (CS) model that allows for 

negative covariance (Littell et al., 2006). 

yijk = μi

 + lj + rk(j) + εijk Model 3

lj

 ~ iid N(0,σ2

Site), rk(j) ~ iid N(0,σ2Rep)

In the CS model, the residual variance structure for the ijth entry by site combination is defined as: 



























1

...

...

...

...

...

...

1

...

1

( ) 2 









 V  ij  CS

where εij = [εij1 εij2 ... εijK] is defined as the vector of residuals for the ith entry at the jth site, σ2

CS denotes the residual 

variance in the CS structure and  denotes the correlation between residuals for the same entry at the same site,  = 

Cov(εijk, εijk’)/σ2

CS. 

REML estimation procedure was used to generate estimates of variance and covariance components, 

and entry means (i.e. LS-Means) across sites were then estimated with standard errors. 
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Model Considering Lower Limit of Quantification (LLOQ)

For the analytes copper, glucoalyssin, gluconapoleiferin, glucoraphanin, and neoglucobrassicin, some 

sample values were below the assay LLOQ. Sample results below the LLOQ were treated as leftcensored observations at the respective assay LLOQ value. Data were analyzed using the following 

linear mixed model: 

yijk = μi

 + lj + rk(j) + (μl)ij + εijk Model 4 

lj

 ~ iid N(0, σ2

Site), rk(j) ~ iid N(0,σ2Rep), (μl)ij ~ iid N(0, σ2Entry×Site), εijk ~ iid N(0, σ2Error)

Where μi denotes the mean of the ith entry (fixed effect), lj denotes the effect of the jth site (random effect), rk(j) denotes 

the effect of the kth block within the jth site (random effect), (μl)ij denotes the interaction between the entries and sites 

(random effect) and εijk denotes the residual for the observation obtained from the plot assigned to the ith entry in the 

k

th block of the jth site. 

The conditional likelihood for each observation, given the random effects, was formulated according to 

the status of the observation (i.e. observed or left-censored) (Thiébaut and Jacqmin-Gadda, 2004): 















    

  



left censored

c

P y c

observed y P y y

f y

ijk

ijk

ijk ijk

ijk ijk

ijk

( ) ( )

( ) ( )

( | )

*

*







 

 

Where  denotes the vector of all random effects, ijk denotes the mean of yijk, denotes the standard normal density 

function, denotes the standard normal cumulative distribution function, y*

ijk denotes the observed sample value of 

yijk, and c denotes the assay LLOQ value.

The conditional likelihood function is a product of all individual conditional likelihoods, and the marginal 

likelihood function is formed when the conditional likelihood function is integrated over all random effects. 

Maximum likelihood (ML) procedure was then used to generate estimates of variance components and 

entry means (i.e. LS-Means) across sites. 

2.2.2. Least Squares Mean (LS-Mean) and Confidence Interval (CI) Calculations 

For each agronomic characteristic or composition analyte, the LS-mean value across sites was estimated 

from the corresponding statistical model for herbicide-treated 73496 canola and the control canola. The 

95% CI for each of the entry means was formed by: 

LS-Mean ± t0.975,v × standard error of LS-Mean 

where t0.975,v denotes the upper 0.025 percentage point of a t-distribution with v degrees of freedom. The degrees of 

freedom were determined by Kenward-Roger method (Kenward and Roger, 1997). 

2.2.3. Statistical Comparisons and Interpretations 

The first step in the evaluation was to test for differences in LS-Mean values between herbicide-treated 

73496 canola and the control canola. The FDR method of Benjamini and Hochberg (Benjamini and 

Hochberg, 1995; Westfall et al., 1999) was applied as a post-hoc procedure to account for multiple 

comparisons due to multiple endpoints (i.e. agronomic characteristics or composition analytes) and 

p-values were adjusted accordingly. Significant differences were established if an adjusted p-value 

<0.05. 

For nutrient composition analytes where a statistically significant difference (adjusted p-value <0.05) was 

identified in the across sites analysis, the respective range of individual values was compared to a 
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tolerance interval. Tolerance intervals containing 99% of the values for corresponding analytes of the 

conventional canola population with 95% confidence level (Graybill, 1976) were derived from data 

collected from seven conventional (i.e. non-modified) commercial canola lines grown at five field locations 

in canola-growing areas of the U.S. and Canada over two growing seasons. The selected canola lines 

represent non-modified canola population with a history of safe use, and the selected environments (i.e. 

site and year combinations) represent canola growth under a wide range of environmental conditions (i.e. 

soil type, temperature, precipitation, and irrigation) and canola maturity group zones. 

2.3. Special Considerations 

 

2.3.1. Agronomic Data Considerations 

Plant Height

Plant height was measured on 10 plants per plot. Plot average (i.e. average of 10 plants) was treated as 

the response for the across sites analysis. 

2.3.2. Nutrient Composition Data Considerations 

Transformation

A natural logarithmic “ln(y)” transformation was performed for the raw data of all nutrient composition 

analytes before statistical analyses. For each analyte, residuals from the across sites analysis were 

examined for validation of the normality and constant variance assumptions. The assumptions were 

reasonably satisfied for most analytes after “ln(y)” transformation. 

For the analyte tryptophan, residual distribution skewed to the left and therefore, a square “(y)2

” 

transformation was performed to the raw data instead. The model assumptions were reasonably satisfied 

for this analyte after “(y)2

” transformation. 

The statistical comparisons were conducted based on the transformed data. The estimated mean values 

and the confidence limits were then back-transformed to the original data scale for reporting purposes. 

Fatty Acids

For some fatty acids analytes, absolute sample values were detected below the assay LLOQ. When 

sample values were converted from absolute values to relative proportions (i.e. percent of total fatty 

acids), sample results that were below the LLOQ were assigned a zero value to reflect a negligible 

proportion. However, these zeros were not “true” zeros, but were some unknown small positive values. 

Even though the origination of these zero values was the LLOQ values, it was not straight-forward to 

determine an LLOQ value on the relative proportion scale for each analyte. Therefore, Model 4 was not 

used to analyze fatty acids analytes with zero values; instead it was deemed appropriate to treat these 

zero sample values as missing values and analyze the non-missing data using Model 2 or Model 3. 

Partial LLOQ Sample Values

For a given analyte (including non-fatty acids and fatty acid analytes), the number of samples below the 

assay LLOQ value determined whether a statistical analysis was conducted. The following rules were 

implemented. 

 If < 80% of samples for each entry were below the LLOQ, then statistical analysis was conducted 

using the appropriate mixed model. 

 If ≥ 80% of samples for a single entry within the study were below the LLOQ, then mixed model 

across sites analysis was not conducted. However, if the numbers of samples below the LLOQ 

were not even between the control canola and herbicide-treated 73496 canola, such observation 
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would be discussed. Descriptive statistics (i.e. arithmetic means and ranges) were calculated for 

analytes that were not statistically analyzed using mixed model analysis. 

2.4. SAS Procedures 

SAS PROC GLIMMIX was used to fit Model 1, SAS PROC MIXED was used to fit Model 2 and Model 3, 

and SAS PROC NLMIXED was used to fit Model 4. All three procedures generated LS-Means and 95% 

confidence intervals, and provided statistical comparisons (i.e. p-values). SAS PROC MULTTEST was 

used to provide adjusted p-values. 
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Appendix 3. Materials and Methods for Determination of GAT4621 Protein 

Concentrations 

Plant tissue concentrations of the GAT4621 protein were determined using enzyme-linked 

immunosorbent assays (ELISA) developed at Pioneer Hi-Bred International, Inc., Johnston, IA, USA. 

3.1. Plant Material 

73496 canola plants of the F1*4 generation (refer to Section III-A, Figure 2), plus near-isoline control 

canola plants, were grown concurrently at six field trial locations near: Portage la Prairie, MB; Saskatoon, 

SK (grain samples not included from this site); Minto, MB; Elm Creek, MB; Ephrata, WA; and Velva, ND. 

Trials were planted in a randomized complete block design containing four blocks per treatment and were 

managed according to standard local agronomic practices, with the exception that the 73496 canola 

plants were treated with two applications of glyphosate herbicide (Touchdown HiTech®), at the first true 

leaf stage and the five-leaf stage, respectively. Plant tissue samples were collected from impartially 

selected, healthy individual plants from each block at the following growth stages: 

 BBCH15 – five true leaves unfolded (whole plant sample) 

 BBCH33 – three visibly extended internodes (whole plant sample) 

 BBCH65 – full flowering; 50% of flowers open on main raceme, older petals falling (whole plant 

and root samples) 

 BBCH90 – senescence (seed sample) 

3.2. Storage and Processing of ELISA samples 

After collection, tissue samples were shipped overnight on dry ice to Pioneer Hi-Bred International, Inc. 

Whole plant samples, which comprised the entire above-ground portion of the plant, were coarsely 

homogenized and all samples (whole plant, root, and seed) were lyophilized, and then finely 

homogenized and stored frozen (≤ -10oC). 

3.3. Protein Extraction from Ground Canola Tissues 

Lyophilized, ground tissue samples were weighed into 1.2-ml tubes at the following target weights: 15 mg 

for whole plant, and 20 mg for root and seed. Each sample was extracted with 0.6 ml of chilled H5 buffer 

solution (90 mM HEPES pH 7.6, 140 mM NaCl, 1% PVP-40, 0.3% Tween-20, 1% bovine serum albumin, 

1% polyethylene glycol, and 0.007% Thimerosal), centrifuged, and the supernatants removed, diluted, 

and analyzed. 

3.4. Determination of GAT4621 Protein Concentrations

Sample extracts were quantitatively analyzed for GAT4621 by sequential double-antibody sandwich 

ELISA using antibodies specific for the GAT protein. Standards (analyzed in triplicate wells) and diluted 

sample extracts (analyzed in duplicate wells) were incubated in a plate pre-coated with GAT-specific 

antibody. Following incubation, unbound substances were washed from the plate. A different GATspecific antibody conjugated to the enzyme horseradish peroxidase (HRP) was added to the plate and 

incubated. Unbound substances were washed from the plate leaving the bound GAT4621 protein 

“sandwiched” between the antibody coated on the plate and the antibody-HRP conjugate. Detection of 

the bound GAT4621-antibody complex was accomplished by the addition of substrate, which generated a 

colored product in the presence of HRP. The reaction was stopped with an acid solution and the optical 

density (OD) of each well was determined using a plate reader. An average of the results from duplicate 

wells was used to determine the concentration of the GAT4621 protein in ng/mg sample weight. 
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SoftMaxc Pro software (Molecular Devices Corporation, Sunnyvale, CA) was used to perform the 

calculations required to convert the OD values obtained by the microtiter plate reader to tissue protein 

concentrations. A standard curve was included on each ELISA plate. The equation for the standard curve 

was generated by the software, which used a quadratic fit to relate the mean OD values to the respective 

standard concentrations (ng/ml). 

Lower limits of quantification (LLOQ) by ELISA were established with each sample matrix and were 0.29 

ng/mg dry weight for whole plant samples, and 0.22 ng/mg dry weight for root and seed samples. For 

each plant tissue type and growth stage, levels of GAT4621 protein in samples from control plants were 

below the LLOQ. 

 

c

 Registered trademark of Molecular Devices Corporation 
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Appendix 4. Characterization of the Plant-Expressed and Microbial-Expressed GAT4621 

Proteins 

4.1. Characterization of the Plant-Expressed GAT4621 Protein 

Physicochemical characterization of the GAT4621 protein isolated from 73496 canola was conducted in 

order to demonstrate equivalence to the microbial-expressed form of the protein used in acute toxicity 

testing and digestibility studies previously conducted for maize event DP-Ø9814Ø-6. This 

characterization of in planta expressed GAT4621 protein from 73496 canola included: molecular weight 

and immunochemical cross-reactivity by SDS-PAGE and western immunoblot analysis; N-terminal amino 

acid sequencing; and tryptic peptide mapping by MALDI-MS. In addition, the GAT4621 protein from 

73496 canola was analyzed for glycosylation as part of the weight of evidence for assessing the potential 

allergenicity. 

In the case of the N-terminal amino acid sequencing and tryptic peptide mapping by MALDI-MS, data 

obtained previously for the microbial-expressed GAT4621 were used for comparison to the 

characterization of the GAT4621 protein from 73496 canola. 

Utilizing these analyses, the equivalency of GAT4621 protein expressed in E. coli to the protein 

expressed in 73496 canola was demonstrated. Therefore, the GAT4621 protein derived from the 

microbial expression system was appropriate for utilization in safety assessment studies as a proxy for 

the GAT4621 protein expressed in 73496 canola. In addition, the GAT4621 protein produced in 73496 

canola is not glycosylated and supports the weight of evidence assessment that the GAT4621 protein is 

unlikely to be an allergen. 

4.2. SDS-PAGE Analysis of the Plant-Expressed GAT4621 Protein 

Analysis of GAT4621 protein isolated from 73496 canola by SDS-PAGE followed by staining revealed a 

prominent band with a relative mobility consistent with the GAT4621 molecular weight of 16.5 kDa 

(Figure 4.1). 

Based on this analysis, the approximate size of the plant-expressed GAT4621 protein was consistent with 

the expected theoretical size of the GAT4621 protein. 

Figure 4.1. SDS-PAGE Analysis of the Plant-Expressed GAT4621 Protein 

A sample of the purified GAT4621 protein (~ 1 μg total protein) from 73496 canola (lane 2) was analyzed by 

SDS-PAGE on 10–20% gradient gels followed by staining with GelCode Coomassie Blue reagent. Molecular 

weight markers (PageRuler Prestained Protein Ladder) were included in lane 1. The major band corresponding 

to the intact size of the GAT4621 protein is indicated. 
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4.3. Western Immunoblot Analysis of the Plant-Expressed and Microbial-Expressed GAT4621 

 Proteins

Western blot analysis of the GAT4621 protein prepared from 73496 canola revealed a predominant 

immunoreactive band that co-migrated with the microbial-expressed GAT4621 and corresponded to the 

molecular weight of 16.5 kDa for the GAT4621 protein (Figure 4.2). 

Based on this analysis, the microbial- and the plant-expressed GAT4621 proteins were determined to be 

equivalent in size and immunoreactivity. 

Figure 4.2. Western Immunoblot Analysis of the Plant-Expressed and Microbial-Expressed 

GAT4621 Proteins 

A sample of the GAT4621 protein (~ 10 ng protein estimated) from 73496 canola (lane 2) and a sample of the 

microbial-expressed GAT4621 protein (Lot: PCF-0005; ~ 15 ng of purified GAT4621; lane 3) were analyzed by SDSPAGE followed by electroblotting onto polyvinylidene difluoride (PVDF) membrane. The blots were probed 

sequentially with mouse anti-GAT monoclonal antibody and horseradish peroxidase-conjugated anti-mouse IgG. 

Blots were then incubated in the presence of a chemiluminescent substrate and the signal was detected with Fujifilm 

Luminescent Image Analyzer. Molecular weight markers (PageRuler Prestained Protein Ladder) were included in 

lane 1. Note: The total protein extract from 73496 canola was loaded in lane 2 and ~ 15 ng of the purified microbialexpressed GAT4621 protein was loaded in lane 3; therefore, the amounts of the GAT4621 protein in lanes 2 and 3 

were not equivalent and the amount of plant-expressed protein was estimated from the blot image. 

4.4. N-Terminal Amino Acid Sequence Analysis of the Plant-Expressed and Microbial-

 Expressed GAT4621 Proteins 

The N-terminal sequence analysis of the GAT4621 protein derived from 73496 canola indicated that the 

primary sequences matched residues 2–14 of the deduced N-terminal sequence of GAT4621 and also 

matched residues 2-11 of the microbial-expressed protein (Figure 4.3). In the plant-expressed GAT4621, 

the lysine signal at position 6 and the aspartic acid signal at position 12, relative to the predicted 

sequence, were low. The N-terminal methionine residue was not detected, which was consistent with 

results from MALDI-MS (Applied Biosystems/MDS SCIEX 4800 MALDI-TOF/TOF) tryptic peptide 

mapping. 

Edman sequencing analysis confirmed that the N-terminal sequences of the 73496 canola-derived 

GAT4621 protein matched both the theoretical protein sequence as well as the microbial-expressed 

sequences. These results provided additional evidence that the plant- and microbial-expressed GAT4621 

proteins are equivalent. 
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73496 Canola 

Figure 4.3. N-Terminal Amino Acid Sequence Analysis of the Plant-Expressed and MicrobialExpressed GAT4621 Proteins 

A sample of purified GAT4621 protein from 73496 canola was analyzed by SDS-PAGE followed by 

electroblotting onto PVDF membrane. The band corresponding to the GAT4621 protein was excised and 

subjected to Edman N-terminal sequencing. For the 73496 canola-derived GAT4621 protein, the aspartic acid 

residue in parentheses at position 12 was assigned based on a low signal. As indicated by Xa

, position 6 was 

not assigned due to a low signal. The theoretical and previously determined N-terminal amino acid sequence for 

microbial-expressed GAT4621 protein are shown for comparison. 

4.5. MALDI-MS Identification of Tryptic Peptides 

MALDI-MS analysis of the tryptic peptides for GAT4621 isolated from 73496 canola identified 25 unique 

peptides. Some of the identified peptides had overlapping sequences and several peptides had a 

cysteine residue modified by propionamide or a methionine residue modified by oxidation. Overall, the 

identified peptides accounted for 88.4% (130/147) of the deduced protein sequence for the GAT4621 

protein derived from 73496 canola (Figure 4.4). In comparison, 76% of the theoretical protein sequence 

was represented by uniquely identified tryptic peptides derived from the microbial-expressed GAT4621 

protein. The N-terminal methionine residue was not identified in matching tryptic peptides derived from 

either plant- or microbial-expressed GAT4621, consistent with the results from N-terminal amino acid 

sequencing. 

As with the N-terminal sequencing, the analysis of tryptic peptides provides additional evidence of 

equivalency between the plant- and microbial-expressed GAT4621 proteins. 

Figure 4.4. Deduced GAT4621 Amino Acid Sequence Showing Matching Peptides from MALDI-MS 

Sequence shown in red correspond to matching tryptic peptides identified for the GAT4621 protein isolated from 

73496 canola. Sequence shown in red with shading correspond to matching tryptic peptides identified for the 

microbial-expressed GAT4621 protein. 

4.6. Protein Glycosylation Analysis of the Plant-Expressed GAT4621 Proteins 

In order to analyze glycosylation of the plant-expressed GAT4621 protein as part of the weight of 

evidence for protein allergenicity, the GAT4621 protein isolated from 73496 canola was analyzed using a 

glycoprotein staining procedure. Plant-expressed GAT4621 protein did not exhibit positive staining with 

the glycoprotein staining reagent (Figure 4.5, lane 4, Glycoprotein staining panel), while the positive 

control horseradish peroxidase was stained and clearly visible (Figure 4.5., lane 3, Total protein staining 

panel). 
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Based on this analysis, the 73496 canola-derived GAT4621 protein was determined not to be 

glycosylated and provided support to the weight of evidence assessment that the GAT4621 protein is 

unlikely to be an allergen. 

Figure 4.5. Glycosylation Analysis of the Plant-Expressed GAT4621 Protein 

Samples containing soybean trypsin inhibitor ( ~ 1 μg) as a negative control (lane 2), horseradish peroxidase (~ 1 

μg) as a positive control (lane 3), and GAT4621 protein ( ~ 1 μg) isolated from 73496 canola (lane 4) were 

subjected to SDS-PAGE and analyzed for the presence of carbohydrates using a periodate-acid-Schiff 

glycoprotein staining reagent (Glycoprotein staining panel). Following image capture, the same gel was stained 

with Coomassie Blue to visual all proteins (Total protein staining panel). Molecular weight markers (PageRuler 

Prestained Protein Ladder) were included in lane 1. 

4.7. Methods for Isolation and Purification of the GAT4621 Protein from Canola Leaf Tissue

The GAT4621 protein was isolated from leaf tissue of greenhouse-grown 73496 canola plants. Each 

plant was tested to confirm the presence of the introduced trait by event-specific PCR. Leaves were 

harvested from plants at the five- to six-leaf stage [BBCH 15-16] and stored frozen (-80oC) until use. 

Approximately 80 g of leaf tissue derived from 73496 canola was extracted by homogenizing with a 

Waring blender in 300 ml Buffer A (50 mM HEPES, pH 7.8, 50 mM NaCl, 1 mM EDTA, 5% glycerol, 

0.007% thimerosal) containing Complete Protease Inhibitors (Roche Applied Science, Indianapolis, IN). 

The homogenate was filtered through four layers of cheese cloth and then centrifuged for 20 minutes at 

approximately 30,500 g. The clarified extract was subjected to 85% saturated ammonium sulfate 

precipitation. The precipitated proteins were resuspended in approximately 60 ml of Buffer A. This sample 

was then desalted with 10DG columns (Bio-Rad Laboratories, Hercules, CA) and pre-equilibrated with 

Buffer B (50 mM MES, pH 7.2, 5% glycerol, 1 mM EDTA). The desalted sample was loaded to an SP 

Sepharose Fast Flow (GE Healthcare Biosciences, Piscataway, NJ) column (10 ml) equilibrated with 

Buffer B. The column was washed with the same buffer (approximately 50 ml) and then the bound 

proteins were eluted with a salt gradient (approximately 100 ml 0-0.5 M NaCl). The GAT4621 fractions 

were collected, pooled and diluted (1:1) with Buffer A. GAT4621 protein was further purified with an 

immuno-affinity column (approximately 3 ml) prepared with GAT-specific mouse monoclonal antibody 

(12C9). The column was sequentially washed with 50 ml each of Buffer A, Buffer C (Buffer A containing 

100 mM NaCl and 0.2% Triton X-100) and Buffer D (Buffer A containing 100 mM NaCl and 0.4% Triton X100). The bound GAT4621 protein was eluted using ImmunoPure IgG Elution Buffer (Thermo Scientific 

Inc., Rockford, IL). Fractions were collected and the protein concentration was estimated by a modified 

Bradford assay (Coomassie Plus Protein Assay Reagent, Thermo Scientific, Inc.). The collected fractions 

were analyzed by SDS-PAGE and western blotting with a GAT-specific antibody (18F9) to confirm the 
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presence of the GAT4621 protein. The fractions containing GAT4621 were pooled and concentrated 

using Nanosep 3K concentrators (Pall Corporation, Ann Arbor, MI). 

4.8. Methods for Isolation and Purification of the Microbial-Expressed GAT4621 Protein 

The GAT4621 protein was expressed in E. coli strain BL21(DE3) as a soluble protein and was purified at 

Aldveron, LLC (Fargo, ND) using cation exchange chromatography, anion exchange chromatography and 

hydrophobic interaction chromatography, followed by diafiltration into 0.1 M ammonium bicarbonate, pH 

7.8. The purified protein was then lyophilized. For characterization, the GAT4621 protein was used in 

the following form: 1 mg of the lyophilized powder was dissolved in 1 ml of 100 mM KCl, 10% methanol, 

and 25 mM HEPES, pH 7.2. 

4.9. Methods for SDS-PAGE Analysis 

SDS-PAGE was performed by first mixing GAT4621 protein samples with Laemmli sample buffer (NuSep 

Limited, Frenchs Forest, Australia) containing 100 mM dithiothreitol and heating at 100°C for 

approximately five minutes. The prepared protein samples were loaded into a 10-20% gradient Ready 

Gel Tris-HCl gel (Bio-Rad Laboratories, Inc., Hercules, CA). PageRuler Prestained Protein Ladder 

(Fermentas, Inc., Vilnius, Lithuania) molecular weight markers were loaded into the gel to provide a visual 

estimate of molecular weight. Electrophoresis was conducted using the Ready Gel Cell system (Bio-Rad 

Laboratories, Inc.) with Tris-glycine running buffer (Fermentas, Inc.) at 150 volts for 60 minutes. 

Upon completion of electrophoresis, the gel was removed from the gel cassette and washed three times 

with deionized water for five to 15 minutes each. The gel was then stained for approximately 60 minutes 

with GelCode Coomassie Blue stain reagent (Thermo Scientific Inc., Rockford, IL), and washed with 

deionized water at least four times for approximately ten minutes each or until the background of the gel 

was clear.

4.10. Methods for Western Immunoblot Analysis 

GAT4621 protein samples were run by SDS-PAGE as described in the previous section. The resulting gel 

was soaked in cathode buffer (60 mM Tris, 40 mM CAPS, 0.1% SDS, pH 9.6) for 10-20 minutes. A 

polyvinylidene difluoride (PVDF) membrane (Bio-Rad Laboratories, Inc.) was briefly placed in 100% 

methanol followed by immersion in anode buffer (60 mM Tris, 40 mM CAPS, 15% methanol, pH 9.6) for 

10-15 minutes. A Trans-Blot SD Semi-Dry Electrophoretic Transfer Cell system (Bio-Rad Laboratories, 

Inc.) was used to transfer proteins from the gel to the membrane at approximately 120 mA for 30-60 

minutes. 

Following protein transfer, the membrane was washed four times for five to ten minutes each in Classic 

Buffer (50 mM Tris-HCl, pH 7.0, 500 mM NaCl, 0.5% Tween-20) and then blocked by incubation in 

phosphate-buffered saline solution with Tween-20 (PBST solution: 8.1 mM phosphate buffer, pH 7.4, 137 

mM NaCl, 2.7 mM KCl, and 0.05% Tween-20) containing 5% non-fat dry milk for 30-60 minutes. The 

blocked membrane was washed four times for 5-10 minutes each in Classic Buffer and then incubated for 

approximately 60 minutes with a GAT-specific mouse monoclonal antibody (18F9) diluted 1:8,000 in 

PBST containing 5% non-fat dry milk. The unbound antibody was removed from the membrane with four 

washes of Classic Buffer for 5-10 minutes each. The membrane was then incubated for approximately 60 

minutes with a secondary antibody (anti-mouse IgG HRP conjugate, Promega, Madison, WI) diluted 

1:10,000 in PBST containing 5% non-fat dry milk, followed by four washes with Classic Buffer for 5-10 

minutes each. The washed blot was immersed in phosphate-buffered saline (PBS: 8.1 mM phosphate 

buffer, pH 7.4, 137 mM NaCl, and 2.7 mM KCl) or PBST solution for at least 5 minutes and then 

incubated with SuperSignal West Dura Chemiluminescent Substrate (Thermo Scientific Inc.) according to 

the manufacturer’s instructions. The signal was detected with Fujifilm Luminescent Image Analyzer (LAS3000, Fujifilm, Hanover Park, IL). 
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4.11. Methods for N-Terminal Amino Acid Sequence Analysis 

Following SDS-PAGE of the 73496 canola-derived GAT4621 protein sample, the resulting gel was 

soaked in cathode buffer (60 mM Tris, 40 mM CAPS, 0.1% SDS, pH 9.6) for approximately 20 minutes. A 

PVDF membrane (Millipore Corp., Bedford, MA) was briefly wetted with 100% methanol, followed by 

immersion in anode buffer (60 mM Tris, 40 mM CAPS, 15% methanol, pH 9.6) for 10-15 minutes. A 

Trans-Blot SD Semi-Dry Electrophoretic Transfer Cell system (Bio-Rad Laboratories, Inc.) was used to 

transfer proteins from the gel to the membrane at 120 mA for approximately 60 minutes. Following protein 

transfer, the membrane was then stained with GelCode Coomassie Blue stain reagent (Thermo Scientific 

Inc.) to visualize the GAT4621 protein band. The resulting GAT4621 band was excised and shipped to 

the Keck Biotechnology Resource Laboratory (Yale University, New Haven, CT) for Edman N-terminal 

amino acid sequencing using the Procise 494 cLC analyzer (Applied Biosystems, Inc., Foster City, CA) 

equipped with an online high performance liquid chromatography system. 

The microbial-expressed GAT4621 protein sample was sent to Crop Genetics (E. I. du Pont de Nemours 

and Company, Wilmington, DE) in 25mM HEPES, pH 7.2, 100mM KCl, 10% methanol. The protein was 

transferred to PVDF membrane using ProSorb Inserts (Applied Biosystems, Foster City, CA) and washed 

with 200 μl 0.1% trifluoroacetic acid in water. The membrane was removed and used for Edman Nterminal amino acid sequencing with the Procise 494 LC analyzer (Applied Biosystems) equipped with an 

online high performance liquid chromatography system. 

4.12. Methods for MALDI-MS Identification of Tryptic Peptides 

Following SDS-PAGE, the 73496 canola-derived GAT4621 protein band was visualized by staining with 

Coomassie Blue and the band was then excised from the gel. The gel slice containing the protein was 

placed in a labeled tube and shipped on dry ice via overnight delivery to the Keck Biotechnology 

Resource Laboratory (Yale University). The protein in the gel slice was digested with trypsin for 18 hours 

at 37°C. An aliquot of the digest was analyzed by matrix assisted laser desorption ionization mass 

spectrometry (MALDI-MS) on an Applied Biosystems/MDS SCIEX 4800 MALDI TOF/TOF Analyzer in the 

reflectron mode of operation. Detected peptide peaks were considered a match if the observed 

experimental mass was within 50 parts per million (ppm) of the theoretical mass of peptides determined 

via in silico trypsin cleavage of the protein sequence. The peptide mass matching was performed using 

GPMAW 7.1. 

Allowances were made for the following potential modifications to the peptides: oxidation of methionine 

(observed value is 15.995 Da greater than the theoretical value) and modification of cysteine residues by 

propionamide adduct during SDS-PAGE (observed value is 71.037 Da greater than the theoretical value). 

For the microbial-expressed GAT4621, the procedures for excising the protein and trypsin digestion were 

similar to those described for the 73496 canola-derived protein. The analysis was conducted on an 

Applied Biosystems/MDS SCIEX 4700 MALDI-Tof-Tof instrument (Applied Biosystems) in the reflectron 

mode of operation. Detected peptide peaks were considered a match if they were within 100 parts per 

million (ppm) of the theoretical mass of peptides determined via in silico trypsin cleavage of the protein 

sequence. 

Allowances were made for the following potential modifications to the peptides: oxidation of methionine or 

tryptophan residues (observed value is 15.995 Da greater than the theoretical value) and modification of 

cysteine residues by acrylamide free radicals during SDS-PAGE (observed value is 71.037 Da greater 

than the theoretical value). 
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4.13. Methods for Protein Glycosylation Analysis 

A GelCode glycoprotein staining kit (Thermo Scientific Inc.) was used according to the manufacturer’s 

instructions to determine whether the GAT4621 protein was glycosylated. The 73496 canola-derived 

GAT4621 protein, a positive control protein (horseradish peroxidase) and a negative control protein 

(soybean trypsin inhibitor) were run by SDS-PAGE as described in the SDS-PAGE analysis section. 

Following electrophoresis, the gel was fixed with 50% methanol for 30 minutes and washed with 3% 

acetic acid. The gel was then incubated with the GelCode oxidizing solution for 15 minutes and washed 

three times with 3% acetic acid. The gel was next incubated with the GelCode glycoprotein staining 

reagent (Thermo Scientific Inc.) for 15 minutes and then treated with the GelCode reducing reagent, 

followed by extensive washing with 3% acetic acid and deionized water. Glycoproteins were detected as 

magenta colored bands on the gel. Following glycoprotein detection, the gel was scanned and the image 

was captured electronically. The same gel was then stained with Coomassie Blue as described previously 

to visualize the total protein content of all protein bands, and an image of the gel was captured 

electronically.
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Appendix 5. Ecological Observations for 73496 Canola

Key to “Range of severity” in Tables 5.1 through 5.4: 

mild – very little disease or insect injury (<10%) visible; 

moderate – noticeable plant tissue damage (10% to 30%); 

severe – significant plant tissue damage (>30%). 

Table 5.1. United States: Insect Stressor Comparison between 73496 Canola and Controls 

Year Permit 

Number State County Insect Stressor Range of Severitya 

Difference 

with 

Control? 

2007 07-255-103n CA Imperial 

Flea beetle (Phyllothreta 

cruciferae or Phyllothreta striolata) 

Mild to 

Moderate No 

Green peach aphid (Myzus 

persicae) 

Mild to 

Moderate No 

2008 08-261-103n CA Imperial 

Flea beetle (Phyllothreta 

cruciferae or Phyllothreta striolata) Mild No 

Green peach aphid (Myzus 

persicae) Mild No 

Aphid (Brevicoryne brassicae) Mild No 

2009 09-016-106n 

ND McHenry Flea beetle (Phyllothreta 

cruciferae or Phyllothreta striolata) Mild No 

WA Grant Imported cabbage moth (Pieris 

brassicae) Mild No 

2009 09-245-104n CA Imperial 

Flea beetle (Phyllothreta 

cruciferae or Phyllothreta striolata) Mild No 

Green peach aphid (Myzus 

persicae) 

Mild to 

Moderate No 

2010 10-020-103n ND Cass 

Grasshopper (Melanoplus 

sanguinipes)

Mild to 

Moderate No 

Aphid (Brevicoryne brassicae) Moderate No 

Diamond back moth 

(Plutella xylostella)

Moderate No 

White cabbage butterflies (Pieris 

rapae)

Mild to 

Moderate No 

White cabbage moths (Mamestra 

brassicae) Mild No 

White cabbage butterflies (Pieris 

rapae) Mild No 
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Table 5.1. United States: Insect Stressor Comparison between 73496 Canola and Controls 

(continued) 

Year Permit 

Number State County Insect Stressor 

Range 

of 

Severitya 

Difference 

with 

Control? 

2010 10-020-103n 

ND 

Ward Flea beetle (Phyllothreta 

cruciferae or Phyllothreta striolata) Mild No 

Grand 

Forks 

Flea beetle (Phyllothreta 

cruciferae or Phyllothreta striolata) Mild No 

Diamond back moth 

(Plutella xylostella) Mild No 

Aphid (Brevicoryne brassicae) Mild No 

McHenry Flea beetle (Phyllothreta 

cruciferae or Phyllothreta striolata) Mild No 

WA Grant Green peach aphid (Myzus 

persicae) Mild No 

a

 Range of severity scores are as follows: Mild – very little insect injury (<10%) visible; Moderate – noticeable plant 

tissue damage (10% to 30%); Severe – significant plant tissue damage (>30%) 

Table 5.2. United States: Disease Stressor Comparison between 73496 Canola and Controls 

Year Permit 

Number State County Disease Stressor Range of 

Severitya 

Difference 

with 

Control? 

2007 07-255-103n CA Imperial None reported N/A N/A 

2008 08-261-103n CA Imperial Downey mildew (Peronospora 

parasitica) Mild No 

2009 09-016-106n 

ND McHenry None reported N/A N/A 

WA Grant Powdery mildew (Erysiphe 

polygoni) Mild No 

2009 09-245-104n CA Imperial None reported N/A N/A 

2010 10-020-103n 

ND 

Cass White mold/Sclerotinia 

(Sclerotinia sclerotiorum)

Mild to 

Moderate No 

Ward None reported N/A N/A 

Grand 

Forks 

White mold/Sclerotinia 

(Sclerotinia sclerotiorum) Mild No 

Fusarium wilt (Fusarium 

oxysporum) Mild No 

Alternaria (Alternaria brassicae) Mild No 

McHenry None reported N/A N/A 

WA Grant Powdery mildew (Erysiphe 

polygoni) Mild No 

a

 Range of severity scores are as follows: Mild – very little disease injury (<10%) visible; Moderate – noticeable plant 

tissue damage (10% to 30%); Severe – significant plant tissue damage (>30%)
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Table 5.3. Canada: Insect Stressor Comparison between 73496 Canola and Controls 

Yeara

 Province Nearest City Insect Stressors Range of 

Severityb 

Difference 

with 

Control? 

2008 

MB 

Morden 

Flea beetle (Phyllothreta cruciferae or 

Phyllothreta striolata) Mild No 

Lygus bug (Lygus sp.) Mild No 

Rosebank 

Flea beetle (Phyllothreta cruciferae or 

Phyllothreta striolata) Mild No 

Lygus bug (Lygus sp.) Mild No 

Crystal City Flea beetle (Phyllothreta cruciferae or 

Phyllothreta striolata) Mild No 

Carman Flea beetle (Phyllothreta cruciferae or 

Phyllothreta striolata) Mild No 

ON Georgetown 

Flea beetle (Phyllothreta cruciferae or 

Phyllothreta striolata) Mild No 

Imported cabbage moth (Pieris brassicae) Mild No 

Lygus bug (Lygus sp.) Mild No 

2009 

AB 

Fort 

Saskatchewan 

Grasshopper (Melanoplus sanguinipes) Mild to 

Severe No 

Cutworm (Euxoa ochrogaster) Mild to 

Severe No 

Gibbons 

Grasshopper (Melanoplus sanguinipes) Mild to 

Severe No 

Cutworm (Euxoa ochrogaster) Mild to 

Severe No 

Riviere Qui 

Barre 

Flea beetle (Phyllothreta cruciferae or 

Phyllothreta striolata) Mild No 

Thrips (Thrips tabaci) Mild No 

Cutworm (Euxoa ochrogaster) Mild No 

MB 

Minto 

Imported cabbage moth (Pieris brassicae) Mild No 

Aphid (Brevicoryne brassicae) Mild No 

Flea beetle (Phyllothreta cruciferae or 

Phyllothreta striolata) Mild No 

Thrips (Thrips tabaci) Mild No 

Rosebank Flea beetle (Phyllothreta cruciferae or 

Phyllothreta striolata) Mild No 

Portage la 

Prairie 

Flea beetle (Phyllothreta cruciferae or 

Phyllothreta striolata) Mild No 

Wellwood Flea beetle (Phyllothreta cruciferae or 

Phyllothreta striolata) Mild No 

Franklin Flea beetle (Phyllothreta cruciferae or 

Phyllothreta striolata) Mild No 

SK Dundurn Alfalfa looper (Autographa californica) Mild No a

 Canadian permits: 2008: 08-PHI-294-CAN01, 2009: 09-PHI1-294-CAN01 

b Range of severity scores are as follows: Mild – very little insect injury (<10%) visible; Moderate – noticeable plant 

tissue damage (10% to 30%); Severe – significant plant tissue damage (>30%)
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Table 5.4. Canada: Disease Stressor Comparison between 73496 Canola and Controls 

Yeara

 Province Nearest 

City Disease Stressor Range of Severityb 

Difference 

with 

Control? 

2008 MB Morden Sclerotinia (Sclerotinia sclerotiorum) Moderate No Rosebank Sclerotinia (Sclerotinia sclerotiorum) Mild to Moderate No 

ON Georgetown Sclerotinia (Sclerotinia sclerotiorum) Mild No 

2009 MB Minto Sclerotinia (Sclerotinia sclerotiorum) Mild No Franklin Sclerotinia (Sclerotinia sclerotiorum) Mild to Moderate No 

SK Dundurn Sclerotinia (Sclerotinia sclerotiorum) Mild No a

 Canadian permits: 2008: 08-PHI-294-CAN01, 2009: 09-PHI1-294-CAN01 

b

 Range of severity scores are as follows: Mild – very little disease injury (<10%) visible; Moderate – noticeable plant 

tissue damage (10% to 30%); Severe – significant plant tissue damage (>30%)
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Appendix 6. USDA and Canadian Field Trials of 73496 Canola

Table 6.1. USDA Field Trials of 73496 Canolaa,b

Year Permit 

Name 

Permit 

Valid 

Date 

State 

Number of 

Counties 

Where Planted 

Acreage 

2007 07-255-103n 10/1/2007 California 1 0.00594 

2008 08-261-103n 10/28/2008 California 1 0.07128 

2009 09-016-106n 2/20/2009 

Idaho 1 0.05 

Minnesota 1 0.035 

New Jersey 1 0.01 

North Dakota 4 0.19264 

Washington 5 0.15535 

2009 09-245-104nc 9/21/2009 California 1 0.55556 

2010 10-020-103nc 3/2/2010 North Dakota 4 0.50327 

Washington 1 0.304 

2010 10-253-111nc 10/5/2010 California 1 0.3 

a

 Plantings through October 19, 2010 are listed. b

 In USDA final reports, 73496 canola (event DP-Ø73496-4) is called DP-073496-4 or DP-73496-4. 

c

 Final field test report not yet due to USDA. 

Table 6.2. Canada Field Trials of 73496 Canolaa

Year Permit Name 

Permit 

Valid 

Date 

Province 

Number of 

Locations 

Planted 

Acreage 

2008 08-PHI-294-CAN 4/29/2008 

Alberta 2 0.28038 

Manitoba 2 0.70093 

Ontario 1 0.22664 

Saskatchewan 1 0.14019 

2009 09-PHI1-294-CAN01 4/30/2009 

Alberta 4 0.33685 

Manitoba 9 0.85154 

Ontario 2 0.24223 

Saskatchewan 7 0.2479 

2010 10-PHI1-294-CAN 4/29/2010 

Alberta 5 2.44817 

Manitoba 1 0.15381 

Ontario 2 0.40707 

Saskatchewan 1 0.87204 

a

 Plantings through October 19, 2010 are listed. 
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Appendix 7. Survey of Potential Substrates for GAT4621 

In enzyme kinetics, kcat is a measure of the turnover rate or speed of the reaction. The higher the kcat, the 

faster the enzyme reaction. KM is the affinity of the enzyme for a substrate or tightness of binding of the 

substrate to the enzyme. The lower the KM, the greater the affinity of the enzyme for the substrate. A 

kcat/KM ratio is the common way to express the catalytic efficiency of the enzyme. The greater the kcat/KM

ratio, the greater the catalytic efficiency of the enzyme for a given substrate. Specificity of an enzyme can 

be judged by comparing the values of kcat/KM for various enzyme substrates. 

In order to examine the specific catalytic efficiency of the GAT4621 enzyme expressed in 73496 canola, a 

substrate specificity study was done with the microbial GAT4621 protein that had previously been 

determined to be equivalent to plant-derived GAT4621 protein from 73496 canola (Section VI-D and 

Appendix 4). 

A dithiobis-2-nitrobenzoate enzymatic end-point assay was used to detect any enzyme activity in a survey 

of various substrates (Siehl et al., 2005) using assay buffer containing 100 mM KCl to represent 

physiological conditions in the plant cytosol and chloroplast (Cuin et al., 2003; Sanders and Bethke, 

2000). Twenty agrochemicals, 21 amino acids and 11 antibiotics were tested as potential substrates for 

the GAT4621 protein. As expected from a previous survey (Siehl et al., 2005) with a Round 11 GAT 

protein that only differed from GAT4621 in 73496 canola by the absence of an alanine residue in the 

second position of the protein sequence, no significant activity was seen with the majority of the 

substrates. 

Of the substrates surveyed for activity with GAT4621, only five amino acid substrates indicated low but 

measurable enzyme activity (sufficiently above the limit of quantitation of the assay): L-aspartate, 

L-glutamate, L-serine, glycine and L-threonine. The remaining amino acids and substrates surveyed 

(other than glyphosate) produced levels of end product near or below the limit of quantitation of the endpoint assay. 

GAT4621 activity on the five amino acids was further characterized using a continuous 

spectrophotometric assay (Siehl et al., 2005) for characterization of kinetic properties (Table 1). The 

affinity of the GAT4621 enzyme for glycine was so low that KM values could not be accurately estimated. 

Therefore, a kcat/KM ratio could not be calculated. 

Table 7.1. kcat/KM for the GAT4621 Enzyme on Glyphosate and Selected Amino Acids 

Substrate 

kcat/KM

(+/- standard deviation) 

min-1 mM-1 

% kcat/KM of glyphosate 

Glyphosate 1063 (+/- 31.8) 100 

Aspartate 12.1 (+/- 1.57) 1.14 

Glutamate 8.32 (+/- 0.46) 0.78 

Serine 0.57 (+/- 0.05) 0.05 

Threonine 0.60 (+/- 0.01) 0.06 

Glycine NDa ND1

a

 ND – Not able to determine because the KM was too high to estimate. 

The results of the GAT4621 substrate survey confirmed the results of the earlier published substrate 

survey for the GAT enzyme. The GAT4621 enzyme was able to use five amino acids as substrates 

(aspartate, glutamate, serine, threonine and glycine), although inefficiently (~0.05 - 1% of that of 

GAT4621 on glyphosate). Further kinetic characterization indicated that the KM of GAT4621 for glycine 

was too high to estimate (low affinity of the GAT4621 enzyme for glycine as substrate). The level of 
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catalytic efficiency of GAT4621 on aspartate, glutamate, serine, and threonine was about 1%, 0.8%, 0.05 

and 0.06%, respectively, of the activity on glyphosate. 

Due to the low levels of GAT4621 enzyme activity on five amino acids as substrates, the concentrations 

of NAA, NAG, NAS, NAT and NAGly in 73496 and control canola seed was measured in the nutrient 

composition studies (Section VIII). 
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Appendix 8. Safety and History of Consumption of Acetylated Amino Acids 

8.1 Summary 

In addition to glyphosate, the GAT4621 enzyme is known to acetylate five amino acids: aspartate, 

glutamate, threonine, serine, and glycine. Acetylated amino acids are ubiquitous in nature, and are part 

of many biological systems in plants and animals. In addition, acetylated amino acids can be used in 

animal feed applications and industrial applications. A large amount of data has been developed on the 

safety of consumption of NAA, NAG, NAT, NAS, and NAGly. Collectively, these data indicate that 

acetylated amino acids have a history of safe use and are as safe for consumption as table salt. 

8.2 Function and Use of Acetylated Amino Acids 

Acetylated amino acids are naturally occurring substances that have been identified in many biological 

systems. Acetylation of N-terminal amino acids is the most commonly observed posttranslational 

modification of cytosolic proteins (Persson et al., 1985; Polevoda and Sherman, 2002). It has been 

estimated that up to 80% of all cytosolic proteins in mammalian systems are N-acetylated (Brown and 

Roberts, 1976; Driessen et al., 1985). Enzymatic acetylation of amino acids using acetyl-CoA as the 

acetyl donor group can occur either cotranslationally or posttranslationally depending on the biological 

system (Polevoda and Sherman, 2000). Enzymes responsible for intracellular acetylation of amino acids 

(N-acetyltransferases) have been identified in rat, yeast, and other eukaryotic organisms (Lee et al., 

1988; Lee et al., 1989; Mullen et al., 1989; Yamada and Bradshaw, 1991a; Yamada and Bradshaw, 

1991b). The biological role of acetylation of N-terminal amino acids of cytosolic proteins has been 

investigated and evidence indicates that this modification protects proteins from proteolysis by 

intracellular aminopeptidases (Berger et al., 1981; Brown, 1979; Jörnvall, 1975). A variety of additional 

roles for N-acetylation of amino acids in biological systems have been established (Polevoda and 

Sherman, 2002). 

Taking into account the wide distribution and biological roles of acetylation, it is not surprising that a 

number of enzymes responsible for deacetylation of acetylated amino acids (i.e., acylases) have also 

been described. It has long been speculated that enzymatic deacetylation of amino acids is a “general” 

phenomenon in mammals because this reaction has been observed in numerous organs (Neuberger and 

Sanger, 1943). It was later hypothesized that this enzymatic reaction plays a role in the salvage of 

acetylated amino acids formed during the metabolic degradation of N-terminal acetylated proteins (Endo, 

1980; Gade and Brown, 1981). To date, four classes of acylases (Types I – IV) that mediate 

deacetylation of acetylated amino acids have been described in mammalian systems that differ with 

regard to distribution and specificity. 

8.3 History of Safe Use of Acetylated Amino Acids 

Acetylation of proteins is commonly employed in the food industry to alter the solubility, water absorption 

capacity and emulsifying properties of protein concentrates (e.g., El-Adawy, 2000; Ramos and Bora, 

2004). Another well-characterized use of acetylated amino acids is in the livestock industry in cases 

when it is unsuitable to use free amino acids in feed. For example, the quality of soy protein fractions can 

be limited by the concentrations of the essential amino acid L-methionine. This deficiency can be 

overcome by supplementation of diets with free L-methionine; however this can lead to development of 

objectionable odors and flavors from conversion of L-methionine to methional by Strecker degradation 

(Ballance, 1961). Therefore, feed may be supplemented with N-acetyl-L-methionine. In rats fed diets 

produced with soy protein isolates, growth and weight gains were similar regardless of whether they were 

supplemented with L-methionine or N-acetyl-L-methionine (Amos et al., 1975; Boggs et al., 1975). 

Metabolism studies have demonstrated that N-acetyl-14C-L-methionine is readily metabolized to 

L-methionine in rats and in human infants (Boggs, 1978; Stegink et al., 1980; Stegink et al., 1982). 

Similarly, metabolic deacetylation of N-acetyl-L-methionine has been reported in in vitro studies using 

rabbit intestinal epithelial cells (Brachet et al., 1991). While these reports demonstrate that this enzymatic 
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deacetylation occurs within the digestive system, there is also evidence that N-acetyl amino acids are 

deacetylated in other tissues (Yoshida and Lin, 1972). 

Nutritional and metabolic studies with the N-acetyl forms of some amino acids have been conducted in 

humans, rats, and pigs. In most cases, these studies have reported that the N-acetyl form of amino acids 

substitute for the constituent amino acid via metabolic deacetylation. Such results have been reported for 

glutamate (Arnaud et al., 2004; Magnusson et al., 1989; Neuhäuser and Bässler, 1986), phenylalanine 

and tryptophan (du Vigneaud et al., 1934) and threonine (Boggs, 1978). Although specific information is 

not available for aspartate, threonine, serine, or glycine, there is no reason to believe these amino acids 

would not also be biologically available when acetylated. 

8.4 Presence of NAA, NAG, NAT, NAS, and NAGly in Biological Systems 

Plants expressing the GAT4621 enzyme are known to contain increased concentrations of some or all of 

the following five acetylated amino acids: NAA, NAG, NAT, NAS, and NAGly. As described earlier, 

acetylated amino acids are known to be widely present in biological systems and some additional 

information about the cellular location and possible function of these five acetylated amino acids are 

described further below. 

8.4.1 NAA 

NAA is a component of the mammalian central nervous system (CNS). It was first isolated from the 

brains of cats in 1956 (Tallan et al., 1956). Since that report NAA has been found to be a common 

constituent of the CNS of all mammals (reviewed in Moffett et al., 2007) where it is located almost entirely 

within the neurons and produced at high concentrations. It has been reported that NAA is the second 

most abundant free amino acid in the mammalian CNS – second only to the concentrations of free 

glutamate (Miyake et al., 1981; Simmons et al., 1991; Tsai and Coyle, 1995; Urenjak et al., 1993). 

The metabolic pathway by which NAA is synthesized in the mammalian CNS has also been determined. 

It is produced enzymatically by acetylation of L-aspartate using acetyl-CoA as a cofactor. The enzyme 

responsible for this reaction is aspartate-N-acetyltransferase (E.C. 2.3.1.17). It is expressed exclusively 

in neuron mitochondria (Demougeot et al., 2004; Truckenmiller et al., 1985). 

Within the mammalian CNS, NAA is readily metabolized to L-aspartate and free acetate by the enzyme 

N-acetyl-L-aspartate amidohydrolase (EC 3.5.1.15; D’Adamo et al., 1973; Goldstein, 1976). This enzyme 

has also been called acylase II and aspartoacylase in published literature and will hereafter be referred to 

as aspartoacylase. Recent studies have demonstrated that aspartoacylase is expressed by 

oligodendrocytes (the cells responsible for synthesis of myelin in the CNS) but not by neurons or 

astrocytes (Baslow et al., 1999; Madhavarao et al., 2004). 

NAA is the primary source of acetate required for lipid synthesis used for axonal myelination during 

development of the mammalian CNS (D’Adamo et al., 1968; Patel and Clark, 1979). Aspartoacylase 

liberates acetate from NAA which is then used in the biosynthesis of lipids used for myelination of the 

axons of the developing CNS. In fact, it has been reported that the activity of aspartoacylase in the brains 

of developing rats correlates with the time course of myelination of the brain (Bhakoo et al., 2001; Kirmani 

et al., 2002; Kirmani et al., 2003); therefore NAA is critical for proper development of the mammalian 

CNS. 

8.4.2 NAG 

NAG has been identified as a component of a number of organs (liver, small intestine, brain, kidneys, 

spleen and heart) in numerous animal species (rats, mice and other animals) using different analytical 

methods (Alonso et al., 1991; Reichelt and Kvamme, 1967; Shigesada and Tatibana, 1971a), and serves 

as the first intermediate in the biosynthesis of arginine in prokaryotes, lower eukaryotes and plants 

(Caldovic and Tuchman, 2003). Glutamate is the most abundant amino acid in the mammalian brain and 

it is a principal neurotransmitter in the central nervous system (CNS; Al-Sarraf and Philip, 2003; reviewed 

by Meldrum, 2000). 
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Biochemical studies have demonstrated that NAG in mammals is produced enzymatically by Nacetylglutamate synthase (NAGS; E.C. 2.3.1.1), which acetylates glutamate using acetyl-CoA as a 

cofactor (Caldovic and Tuchman, 2003). This occurs in mitochondria of parenchymal cells from most 

tissues, as high levels of NAGS activity and NAGS specific mRNA have been identified in liver and small 

intestinal mucosal cells of rats and humans though neither NAGS protein nor mRNA have been detected 

in brain tissue (Caldovic et al., 2002a; Caldovic et al., 2002b; Uchiyama et al., 1981). 

Mammalian metabolism of NAG is also well characterized. NAG is hydrolyzed enzymatically by 

aminoacylase I (EC 3.5.1.14) to glutamate and acetate within the cytosol of kidney and liver parenchymal 

cells (Reglero et al., 1977; Shigesada and Tatibana, 1971b). The primary biological activity of NAG is the 

allosteric regulation of carbamyl phosphate synthetase I (CPSI; E.C. 6.3.4.16) activity; the first enzymatic 

step in the urea cycle which is responsible for elimination of excess ammonia from metabolic processes 

(Caldovic and Tuchman, 2003). CPSI is an intramitochondrial enzyme that converts ammonia and 

bicarbonate into carbamyl phosphate which is subsequently combined with ornithine via ornithine 

transcarbamylase (EC 2.1.3.3) to form citrulline which is then exported from the mitochondria. The activity 

of CPSI is so dependent on NAG that the activity of this enzyme is virtually undetectable in the absence 

of NAG (Caldovic and Tuchman, 2003; Hall et al., 1958). This dependence has clinical relevance for 

humans and other mammals because deficiency of NAG can arrest urea cycle metabolism and lead to 

hyperammonemia (Caldovic et al., 2002a). 

8.4.3 NAT

NAT is a derivative of the amino acid L-threonine bearing an acetyl group covalently linked to the amine 

nitrogen. It has been identified as the N-terminal amino acid of a number of dietary proteins though the 

biological function is unknown (Polevoda and Sherman, 2003). 

Acetylation of proteins has been reported to protect proteins from degradation (Persson et al., 1985), 

assist in the export of soluble proteins from the cell (Chang et al., 2008), and block activation of signaling 

pathways by antagonizing phosphorylation of threonine residues (Mukherjee et al., 2007). 

8.4.4 NAS

NAS is an acetylated derivative of the amino acid L-serine and has been identified on the N-terminus of 

proteins as a commonly occurring post-translational modification of proteins in eukaryotes (Brown and 

Roberts, 1976; Polevoda and Sherman, 2003). Acetylation of proteins has been reported to protect 

proteins from degradation (Persson et al., 1985), assist in the export of soluble proteins from the cell 

(Chang et al., 2008), and block activation of signaling pathways by antagonizing phosphorylation of serine 

residues (Mukherjee et al., 2007). 

Within nature, NAS is one of the most commonly acetylated amino acids of proteins. In fact, it has been 

estimated that approximately 90% of proteins with N-terminal serine residues are acetylated (Driessen et 

al., 1985). Accordingly, proteins containing NAS are likely to be routinely consumed in the diet from a 

wide variety of plant and animal sources (Brown and Roberts, 1976; Persson et al., 1985). 

Plants and microorganisms are able to reduce inorganic sulfur resulting in L-cysteine biosynthesis. A 

pathway for this has been described in microorganisms involving free NAS. Through a feedback inhibition 

mechanism, L-cysteine inhibits the biosynthesis of O-acetyl-L-serine, which is the precursor for NAS, an 

inducer of the cysteine regulon. (Kredich, 1996). 

8.4.5 NAGly

A number of naturally occurring proteins from eukaryotic cells contain NAGly, including; cytochrome c, 

hemoglobin and ovalbumin (Brown and Roberts, 1976). Specific activity for the hydrolysis of NAGly has 

been observed in liver, kidney and brain tissue (Bray et al., 1949; Bray et al., 1950; Giardina et al., 2000; 

Goldstein, 1976; Mounter et al., 1958; Reglero et al., 1977). 
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Aminoacylase I (ACY1; EC 3.5.1.14) is an enzyme involved in the cytoplasmic degradation of Nacetylated derivatives of serine, glutamic acid, alanine, methionine, glycine, leucine, and valine. When 

aminoacylase activity is compromised, as in the case of ACY1 deficiency, there is marked increase in 

urinary excretion of acetylated amino acids including NAGly (Gerlo et al., 2006; Sass et al., 2006). 

8.5 History of Safe Consumption of NAA, NAG, NAS, NAT, and NAGly 

NAA, NAG, NAS, NAT, and NAGly are normal components of food and feedstuffs. As demonstrated by 

the compositional analysis described in Section VIII and more extensively in submissions for previously 

approved 98140 maize (FDA, 2008; USDA, 2009), these compounds are found in plant species including 

canola, are not novel, and are normal components of animal diets. 

8.6 Toxicology Studies Conducted with NAA, NAG, NAS, NAT, and NAGly 

A substantial number of toxicology studies have been conducted with NAA, NAG, NAS, NAT, and NAGly. 

There was no evidence of mutagenicity in individual in vitro and in vivo studies with NAA, NAG, NAS, 

NAT and NAGly (Harper et al. 2009; Harper et al. 2010; Karaman et al., 2009; van de Mortel et al. 2010a; 

van de Mortel et al. 2010b). In addition, no adverse effects were observed in rats following acute oral 

exposure to NAA, NAG, NAS, NAT and NAGly individually at 2000 mg/kg of body weight or following 

repeated dose dietary exposure to approximately 1000 mg/kg of body weight (Delaney et al., 2008; 

Harper et al. 2009; Harper et al. 2010; van de Mortel et al. 2010a; van de Mortel et al. 2010b). Mortalities 

and clinical signs of toxicity were observed in rats that were orally dosed with NAA at 5000 mg/kg of body 

weight in an acute toxicity study, indicating that the acute toxicity of this particular substance is similar to 

that reported for table salt (Delaney, 2010). In addition, there was no evidence of adverse effects in 

longer term (i.e., 90 day) NAA feeding studies at doses of approximately 500 mg/kg of body weight in 

which brain myelination was evaluated (Karaman et al., 2011). A two generation reproductive toxicity 

study with NAA that included evaluation of brain lipid myelination and tissue concentrations of NAA in 

which no evidence of adverse effects were observed was also conducted, however, this study has not yet 

been published. The full list of toxicology studies conducted by Pioneer is listed in Table 8.1. 

Table 8.1. Full List of Studies Conducted with NAA, NAG, NAS, NAT, and NAGly

Acetylated 

Amino Acid Publication Study 

NAA 

Karaman et al., 2009 Ames in vitro mutagenicity 

Bone marrow micronucleus in vivo mutagenicity 

Delaney et al., 2008 and Delaney, 

2010 Acute oral toxicity 

Delaney et al., 2008 28-Day repeated dose oral toxicity 

Karaman et al., 2011 90-Day repeated dose oral toxicity 

Not yet published 2-Generation reproduction toxicity 

NAG Harper et al., 2009 

Ames in vitro mutagenicity 

Bone marrow micronucleus in vivo mutagenicity 

Acute oral toxicity 

28-Day repeated dose oral toxicity 

NAT van de Mortel et al., 2010a 

Ames in vitro mutagenicity 

Bone marrow micronucleus in vivo mutagenicity 

Acute oral toxicity 

28-Day repeated dose oral toxicity 

NAS van de Mortel et al., 2010b 

Ames in vitro mutagenicity 

Bone marrow micronucleus in vivo mutagenicity 

Acute oral toxicity 

28-Day repeated dose oral toxicity 

NAGly Harper et al., 2010 

Ames in vitro mutagenicity 

Bone marrow micronucleus in vivo mutagenicity 

Acute oral toxicity 

28-Day repeated dose oral toxicity 
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Appendix 9. Herbicide Resistant Weeds 

9.1. Evolution of Herbicide Resistant Weeds 

Weeds will tend to adapt and circumvent any single control mechanism. Instances of herbicide resistant 

weeds have occurred for many herbicide classes including acetyl-CoA carboxylase and acetolactate 

synthase (ALS) inhibitors, glyphosate, hormone, triazines and other photosystem II inhibitors (Table 9.1; 

Heap, 2010b). 

Herbicide resistance usually evolves in only one or two weed species in an area, even though a much 

larger number of weeds are exposed to the same herbicide selection intensity. Nonetheless, weed 

resistance to herbicides currently affects hundreds of thousands of fields and the most widely used 

herbicides (Heap, 2010a). According to a recent survey, more than 348 herbicide resistant weed 

biotypes (representing 194 species) are confirmed to be present in agricultural fields around the world 

(Table 9.1; Heap, 2010b). In this survey, several criteria are used to confirm resistance. Among these 

criteria, a resistant weed must meet certain specific definitions, is confirmed through unbiased 

experimentation, is able to pass the phenotype to subsequent generations, and has a practical economic 

impact to growers (Heap, 2005). Resistant weeds often increase the cost of crop production and limit the 

effectiveness of herbicides that can be used and the crops that can be grown. Growers use a variety of 

approaches to limit the impact of resistant weeds on crop productivity. 

A graph of the number of confirmed herbicide resistant weeds by herbicide class and by year is provided 

in Figure 9.1. Globally, confirmed glyphosate resistant weeds do not represent the largest class of 

herbicide resistant weeds (Figure 9.1). One hundred thirty-two herbicide resistant weed biotypes are 

confirmed in the United States, with 12 of these resistant to the glyphosate (glycine) class of herbicides 

(Heap, 2010c). 

When growers indicate that their "weeds have become resistant," they really mean that the population of 

resistant weed biotypes has increased to an unacceptable level. The spread of a resistance phenotype 

depends primarily upon the exposure or selection applied by the herbicide. When an herbicide is applied, 

most of the susceptible weeds die while the resistant weeds survive, mature, and produce seed. 

Although the resistant population may be small, repeated application of the same herbicide continues to 

increase the proportion of resistant weeds in the population. 

Not all weed shifts are driven by a genetically based biochemical capacity to survive exposure. For 

example, weeds with delayed emergence and slower development are also able to avoid exposure to the 

herbicide (Hilgenfeld et al., 2004). 

Herbicide resistance can become an ecological problem if the resistant weed biotype replaces the nonresistant biotype in the weed population. Even then, the shift to an herbicide resistant population of 

weeds has ecological consequence only if the resistant population cannot be controlled with other 

herbicides or other control practices. This is generally not the case. Many hundreds of cases of resistant 

weeds have been documented worldwide, but resistance is usually not a limiting factor for crop 

production. 

In spite of the evolution of herbicide resistance in weed populations, US growers continue to have many 

herbicides and management options for weed control. Even so, growers must always be concerned 

about herbicide sustainability and the economic consequences of losing any herbicide technology due to 

the evolution of resistant weeds. 
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Table 9.1. Herbicide Resistant Weeds Summary Table (Heap, 2010b) 

Herbicide Group Site of Action Example 

Herbicide 

Total 

Number of 

Biotypes 

ALS inhibitors 

Inhibition of acetolactate synthase 

ALS (acetohydroxyacid synthase 

AHAS) 

Chlorsulfuron 107 

Photosystem II 

inhibitors 

Inhibition of photosynthesis at 

photosystem II Atrazine 68 

ACCase inhibitors Inhibition of acetyl CoA carboxylase 

(ACCase) Diclofop-methyl 37 

Synthetic Auxins Synthetic auxins (action like 

indoleacetic acid) 2,4-D 28 

Bipyridiliums Photosystem-I-electron diversion Paraquat 25 

Ureas and amides Inhibition of photosynthesis at 

photosystem II Chlorotoluron 21 

Glycines Inhibition of EPSP synthase Glyphosate 20 

Dinitroanilines and 

others Microtubule assembly inhibition Trifluralin 10 

Thiocarbamates and 

others 

Inhibition of lipid synthesis - not 

ACCase inhibition Triallate 8 

PPO inhibitors Inhibition of protoporphyrinogen 

oxidase (PPO) Oxyfluorfen 4 

Triazoles, ureas, 

isoxazolidiones 

Bleaching: Inhibition of carotenoid 

biosynthesis (unknown target) Amitrole 4 

Chloroacetamides and 

others 

Inhibition of cell division (Inhibition of 

very long chain fatty acids) Butachlor 4 

Nitriles and others Inhibition of photosynthesis at 

photosystem II Bromoxynil 3 

Carotenoid 

biosynthesis inhibitors 

Bleaching: Inhibition of carotenoid 

biosynthesis at the phytoene 

desaturase step (PDS) 

Flurtamone 2 

Arylaminopropionic 

acids Unknown Flamprop-methyl 2 

4-HPPD inhibitors 

Bleaching: Inhibition of 4-

hydroxyphenyl-pyruvate-dioxygenase 

(4-HPPD) 

Isoxaflutole 1 

Mitosis inhibitors Inhibition of mitosis / microtubule 

polymerization inhibitor Propham 1 

Cellulose inhibitors Inhibition of cell wall (cellulose) 

synthesis Dichlobenil 1 

Organoarsenicals Unknown MSMA 1 

Total number of resistant biotypes 348 
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Figure 9.1. Number of Resistant Biotypes of Weeds by Herbicide Class and Year (Heap, 2010d) 

9.2. Characteristics of Glyphosate 

Glyphosate is a broad spectrum herbicide that was introduced in the 1970s for management of annual, 

perennial and biennial herbaceous grasses, sedges, and broadleaves, as well as woody brush and trees 

(Franz et al., 1997). 

Glyphosate controls plants by inhibiting the enzyme EPSPS (5-enolpyruvylshikimate-3-phosphate 

synthase). EPSPS is an essential enzyme in the shikimate pathway that ultimately leads to the 

production of aromatic amino acids (tryptophan, tyrosine, and phenyIalanine). The shikimate pathway for 

synthesizing aromatic amino acids, and therefore the enzyme EPSPS, is found in plants, bacteria and 

fungi, but not animals. 

The structure of glyphosate resembles the structure of the substrate for EPSPS, which is 

phosphoenolpyruvate (PEP). Therefore, glyphosate competes with PEP for the enzyme’s active site and 

prevents conversion of PEP to the precursor that is required in the synthesis of aromatic amino acids. 

Aromatic amino acids are essential for many plant processes such as protein synthesis, cell wall 

formation, pathogen defense and hormone production. At high rates, glyphosate is toxic to virtually all 

plants. 

9.3. Evolution of Resistance to Glyphosate 

For more than two decades, the evolution of glyphosate resistance was not perceived as a problem 

(Bradshaw et al., 1997). This opinion was based on the difficulty of discovering a fully functional 5-

enolpyruvylshikimate-3-phosphate synthase (EPSPS) that was insensitive to glyphosate, the inability of 

plant species to enzymatically deactivate glyphosate, the lack of soil activity of glyphosate (thus reducing 

the selection pressure due to exposure), and the empirical observation that no resistant weeds had 

appeared after at least 20 years of use. 
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Known theoretical ways for weeds to develop herbicide resistance did not seem likely for glyphosate 

(Jasieniuk, 1995). Over-production of the EPSPS target site did not increase resistance enough for 

plants to survive glyphosate amounts used in agricultural settings (Kishore and Shah, 1988). The EPSPS 

modifications that conferred glyphosate resistance in bacteria were inside the enzyme’s active site, which 

reduced its catalytic efficiency and thus would probably reduce plant fitness (Padgette et al., 1995). No 

higher plants could be found metabolically inactivate glyphosate (Dyer, 1994). 

However, the views about the ability of weeds to evolve glyphosate resistance changed in 1996 when 

glyphosate-resistant Lolium rigidum, was discovered in Australia (Powles et al., 1998; Pratley et al., 

1999). Since then, glyphosate-resistant biotypes in at least 19 other weed species have been confirmed 

globally (Heap, 2010a), and it is estimated that more than several million acres are now affected by 

glyphosate resistant weeds (Owen, 2010). 

Another economically significant glyphosate-resistant weed is horseweed (Conyza canadensis); it was 

confirmed in 2000 as the first annual broadleaf species with glyphosate resistance (Heap, 2010a; 

VanGessel, 2001). Currently, glyphosate-resistant horseweed is estimated to be infesting over several 

million acres in the U.S. across 17 states (Heap, 2010a). 

The molecular basis for weed resistance to glyphosate is not understood for all weeds that have been 

identified, though some mechanisms have been elucidated. Initial studies of various weed species 

revealed EPSPS target site insensitivity (Baerson et al., 2002). Target site-based resistance has been 

identified for some weed species, including Eleusine indica, Lolium multiflorum, and Lolium rigidum

(Powles and Preston, 2006). Differences in translocation and transport to the chloroplast are also 

important in some weeds, such as horseweed (Conzya canadensis) and Lolium rigidum (Feng et al., 

2004; Lorraine-Colwill et al., 2003; Powles and Preston, 2006). Because weed populations can be 

diverse genetically, any particular weed species may evolve different mechanisms of resistance, as is the 

case for Lolium rigidum (Powles and Preston, 2006). Most recently, EPSPS gene duplication and 

increased gene expression has been hypothesized to explain glyphosate resistance in Amaranthus 

palmeri (Gaines et al., 2010). 

The effectiveness, economic benefits, and ease of using glyphosate have led to repeated applications, 

year after year, in areas where glyphosate tolerant biotech crops are grown. The practice of continuously 

planted glyphosate-resistant soybean was implicated in the development of glyphosate-resistant 

horseweed in Delaware (VanGessel, 2001). This intensive use has resulted in a high selection pressure 

for weeds that inherently are difficult to control with glyphosate (Culpepper, 2004). Eventually, this 

selection pressure can lead the spectrum of weeds in the fields to a shift to those weeds that inherently 

can tolerate glyphosate. 

As predicted, spectrum shifts to weed populations with endogenous glyphosate resistance have occurred 

more rapidly than evolved resistance in response to glyphosate exposure (Shaner, 2000). For example, 

in Iowa, common waterhemp (Amaranthus rudis) and velvetleaf (Abutilon theophrasti) became a concern 

in glyphosate-resistant soybean soon after crop commercialization (Owen, 1997). 

9.4. Stewardship of Herbicide Tolerant 73496 Canola 

In order to best manage their weed populations, growers understandably need to know where to obtain 

information on whether herbicide resistant weed biotypes have been reported in their area. It is important 

to understand that when weeds develop resistance to an herbicide, they do so under specific 

circumstances (heavy reliance on a single herbicide mode of action) at a specific location. Weeds that 

develop resistance to herbicides are referred to as “biotypes” to indicate that only a sub-type of the 

species has developed resistance. If the resistant weed biotypes are controlled effectively by mechanical 

means or use of alternative mode-of-action herbicides, the problem may be limited to a specific locality in 

a specific year. 

Growers can obtain information about resistant weed biotypes reported in their area from many sources, 

including their local crop protection chemical dealers, crop protection chemical company representatives, 
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their state Department of Agriculture, University Extension Services, and local crop consultants. The 

most timely and location specific information can likely be provided by their county extension agent, as 

well as their local seed and/or chemistry providers. The USDA Cooperative Extension System offices are 

staffed by one or more experts who provide practical, research-based information to agricultural 

producers and others in rural areas and communities of all sizes. Some of this information is also 

available to growers from the university extension services, such as the University of Minnesota 

Extension (Gunsolus, 2002) and North Dakota State University (http://www.ag.ndsu.edu/weeds/herbicideresistant-weeds). 

The Glyphosate, Weeds, and Crops Website (http://www.glyphosateweedscrops.org/), a website devoted 

to glyphosate stewardship, also develops and compiles member publications and news reports on 

managing glyphosate-resistant weeds. DuPont is one member of this website and membership also 

includes major universities and agricultural institutions. 

The weed science community is also a valuable source of information about resistant weed biotypes that 

have been reported (International Survey of Herbicide Resistant Weeds, www.weedscience.org). They 

also monitor the evolution of herbicide resistant weeds. However, because the information posted on 

their website relies on extension agents and weed scientists to self-report, it may not be completely 

updated with information for the grower's local area.

DuPont’s Pioneer Hi-Bred International, Inc. and Crop Protection Chemicals businesses both have long 

histories of product stewardship. Examples of these efforts are detailed below. 

9.4a. Local Weed Management

In collaboration with university investigators, private consultants, other manufacturers, and growers, 

DuPont Crop Protection field development personnel conduct hundreds of field trials annually to refine 

existing recommendations and to investigate potential new active ingredients and herbicide combinations 

deployed in an Integrated Weed Management context to improve solutions for existing and emerging 

weed problems. Within Pioneer, field agronomists give presentations to local audiences about resistant 

weeds and best management practices. In addition, they assist customers in making crop management 

decisions, including options for managing weeds. 

9.4b. Product Labeling 

For at least 20 years, all DuPont herbicide labels have carried voluntary statements regarding resistant 

weed management (Figure 9.2). These labels actively promote an integrated weed management 

philosophy as seen in the second paragraph: 
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Figure 9.2. Resistant Weed Management Label
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9.4c. Training and Education of Sales Representatives and Agronomists

DuPont Crop Protection: In addition to the formal academic training of technical, marketing, and sales 

professionals, DuPont Crop Protection has a mandatory on-line interactive training course with intensive 

and comprehensive coverage of product stewardship generally and herbicide resistance specifically. This 

training is administered globally to all employees involved in weed management recommendations and is 

also provided to other interested parties as a community education contribution. The technical, 

marketing, and sales professionals are required to be certified in this training by receiving a passing 

grade of 90% or higher in each of the eight modules. A person unfamiliar with this topic is expected to 

take about twelve hours to complete the internal version of the training. This is the first training of its kind 

to be awarded Certified Crop Adviser (CCA) credits by the American Society of Agronomy. The CCA 

standard allows growers, employers, and other organizations to help manage risk by enabling them to 

have assurance that a person has the appropriate professional qualifications. 

Pioneer: The Pioneer sales force, made up of agronomists, account managers and sales 

representatives, and customers receive ongoing mandatory and voluntary stewardship training 

throughout the sales season through a variety of tactics. These include stewardship training sessions 

administered in person or communicated electronically by Pioneer Stewardship personnel. The Pioneer 

sales force also utilizes a proprietary computer software system to access information about products, 

traits, crop management practices, sales transactions and required stewardship documentation. This 

formal training is in addition to the practical experience of the Pioneer sales force working with growers on 

the positioning and use of products containing biotech traits, including herbicide resistant traits. 

9.4d. Technical Bulletins Provided to Seed Customers and the Public 

Direct Mail 

Pioneer’s Growing Point magazine is mailed to all Pioneer customers, prospects and employees. This 

magazine contains information about products, marketing programs, production and management 

practices and advice on agronomic application of traits and technologies. 

Information on Websites

 Pioneer’s “Growing Point” site (https://www.pioneer.com/growingpoint/login/login.jsp) has an 

extensive agronomy section, with access to information about many different crops and crop 

management practices. Technical bulletins such as “Crop Insights” have been published for the past 

15 years, and are a good example of how Pioneer makes the latest information available to growers. 

 The DuPont Biotechnology website (http://www2.dupont.com/Biotechnology/en_US/) contains an indepth “Science Knowledge” section addressing “Herbicide Resistant Crops and Weed Management: 

Scientific Summary and the DuPont Perspective”, with a section devoted to Integrated Weed 

Management, as well as an FAQ section that addresses weed control and Integrated Weed 

Management. There is also a Scientific Summary on Integrated Weed Management and Herbicide 

Resistance: 

(http://www2.dupont.com/Biotechnology/en_US/science_knowledge/herbicide_resistance/moreinfo8.h

tml). 

9.4e. Involvement in Industry Groups 

DuPont Crop Protection is a long-standing participant in the Herbicide Resistance Action Committee 

(HRAC), an industry-based group supported by Croplife International. Their stated mission is to 

“Facilitate the effective management of herbicide resistance by fostering communication and co-operation 

between industry, government and farmers.” They work towards fostering responsible attitudes towards 

herbicide use, communicating herbicide resistance management strategies and support their 

implementation through practical guidelines, and they seek active collaboration with public and private 
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researchers, especially in the areas of problem identification and devising and implementing management 

strategies. 

9.4f. Involvement with Academic Groups 

Pioneer and DuPont personnel interact with academic weed scientists in conducting trials at university 

sites as well as seeking input from them regarding weed management strategies. 

9.4g. Customer Satisfaction and Weed Resistance Management Plan 

Pioneer and DuPont Crop Protection are committed to active market presence wherever we sell our 

products. Consistent with our stewardship principles, all of our business teams are required to maintain 

an active contact with customers and awareness of end-user practices and a capability to respond rapidly 

to issues arising. DuPont Crop Protection maintains a customer satisfaction database to track and 

address any complaints, including for resistant weeds. Field employees are trained and provided tools 

and processes for responding to inquiries regarding product performance (or any potential impacts on 

human health or environment).
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 Appendix 10. GAT4621 New Protein Consultation 
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References 

I. Rationale for the Development of 73496 Canola 

Brookes G, Barfoot P (2010) GM crops: global socio-economic and environmental impacts 1996-2008. 

PG Economics Ltd, Dorchester, UK 

Duke SO, Powles SB (2009) Glyphosate-Resistant Crops and Weeds: Now and in the Future. 

AgBioForum 12: 346-357 

Smyth S, Gusta M, Phillips P, Castle D (2010) Assessing the Economic and Ecological Impacts of 

Herbicide Tolerant Canola in Western Canada. Alberta Canola Producers Commission, 

http://canola.ab.ca/assessing_the_economic_and_ecological_impacts_of_herbicide_tolerant_canola_in_

western_canada.aspx 

II. The Biology of Canola

CFIA (1994) The Biology of Brassica napus L. (Canola/Rapeseed). Canadian Food Inspection Agency, 

Ottawa, Canada, BIO1994-09 

OECD (1997) Consensus document on the biology of Brassica napus L. (oilseed rape). Organisation for 

Economic Cooperation and Development, Paris, OECD/GD(97)63 

III. Method of Development of 73496 and 61061 Canola

Chen W, Tulsieram L, inventors. May 10, 2007. Microprojectile Bombardment Transformation of 

Brassica. United States Patent Application No. 11/270,996 

Klein TM, Wolf ED, Wu R, Sanford JC (1987) High-velocity microprojectiles for delivering nucleic acids 

into living cells. Nature 327: 70-73 

 

IV. Donor Genes and Regulatory Sequences Canola

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Addendum to

Petition for the Determination of Nonregulated Status for

Herbicide Tolerant 73496 Canola

11-063-01p

COMPREHENSIVE REVIEW OF FERAL CANOLA

Submitting Company:

Pioneer Hi-Bred International, Inc.

7100 NW 62nd Avenue

PO Box 1014

Johnston, IA 50131-1014

Submitted by:

Natalie Weber, Registration Manager

Pioneer Hi-Bred International, Inc.

DuPont Agricultural Biotechnology

DuPont Experimental Station, E353/305C

PO Box 80353

Wilmington DE 19880-0353

Telephone: 302-695-8160

Fax: 302-695-3075

Prepared by:

Aimee Hyten, Don MacKenzie, and Tracy Rood

OECD Unique Identifier: DP-Ø73496-4

September 2, 2011

NO CBI
Addendum to 73496 Canola petition Page 2 of 7

11-063-01p September 1, 2011

BRS has requested additional information and a sound justification for why weedy traits of 

73496 canola are neither plant pest nor “issues of concern”. This topic is touched on briefly in 

Section IX-C (“Weediness Potential of 73496 Canola” of the petition, and this addendum 

provides additional comprehensive information on the potential for and plant pest risk of 

establishment of 73496 feral canola populations. The text in this addendum is copied from the 

Environmental Report that Pioneer is submitting to the BRS NEPA team.

Feral Canola 

Feral canola plants are crop-derived plants occurring outside of cultivated areas, most commonly 

in ruderal (i.e., non-cropped, disturbed) habitats such as field margins, along transportation 

corridors (e.g., roadsides, paths, railway lines), near seed/grain handling facilities (e.g., ports, 

storage facilities), and wastelands where they can survive and reproduce without human 

intervention ((Bagavathiannan and Van Acker, 2008). There are a number of ways by which 

feral canola populations can originate including from spillage of seed during its transport to and 

from the field, redistribution of seed by field equipment, seed dispersal by birds and mammals, 

or as a result of bulk seed handling operations.

Canola is a plant of disturbed habitats in that it takes advantage of disturbed sites due to its early 

germination potential and capacity to capture resources quickly. However, canola is a poor 

competitor and lacks the ability to establish stable populations in undisturbed natural habitats 

(Crawley et al., 2001; Crawley et al., 1993; Warwick et al., 2008). Unless the habitat is 

regularly disturbed, or seed replenished from outside, canola will be displaced over time by other 

plants.

As reviewed by Devos et al. (2011a), while many feral canola populations have been found to be 

transient at a local scale, persisting from one to four years (e.g., (Crawley and Brown, 1995; 

Crawley and Brown, 2004) this is likely counterbalanced at a landscape scale by repeated seed 

addition and redistribution. The contribution of seed derived from resident feral plants to the 

persistence of feral canola populations is still a matter of some discussion. A survey conducted 

in central France indicated that local seed input from resident feral plants accounted for less than 

10 percent of subsequent feral populations ((Pivard et al., 2008), while a four-year field survey 

and assessment of genetic variation in feral canola populations in northwest Germany indicated 

that the percentage of feral plants setting seeds ranged between 30−48% (Elling et al., 2009). 

Thus, feral canola may be considered long-lived at a landscape scale, where the high rate of local 

population extinctions is compensated by the establishment of new populations founded by 

repeated fresh seed spills from agricultural fields and transport, with some proportion of 

populations resulting from seed emerging from the feral soil seedbank.

Because of concerns that transgenic herbicide-tolerant canola might be more weedy or invasive, 

or that the introduced genes would be transferred by pollen to wild relatives, whose hybrid 

offspring might be more weedy or invasive, there have been numerous monitoring surveys 

conducted to assess the presence of transgenes in feral canola populations (Table 11). Not 

surprisingly, in regions where there has been widespread adoption of transgenic herbicide-
Addendum to 73496 Canola petition Page 3 of 7

11-063-01p September 1, 2011

tolerant canola over a number of years, such as western Canada and some states in the U.S., the 

prevalence of transgenes in feral canola populations is similar to the proportion of cultivated 

canola that is GE (Knispel et al., 2008; Yoshimura et al., 2006). For example, in a survey of 

feral canola plants conducted in North Dakota, approximately 86% of the feral plants collected 

tested positive for either the glyphosate- or the glufosinate-tolerance trait, and there were two 

instances where these two traits were found combined within single plants (Schafer et al., 2010). 

Likewise in countries that import significant quantities of canola, but where transgenic herbicidetolerant canola is not commercially cultivated, surveys of feral canola populations occurring 

around transportation routes and port areas have confirmed the presence of glyphosate- and 

glufosinate-tolerance traits (Aono et al., 2006; Kawata et al., 2009; Nishizawa et al., 2009; Saji

et al., 2005). In Japan, which imports more than 2 million tonnes of canola annually for 

crushing, the frequency of herbicide-tolerant traits in feral canola populations has been found to 

vary significantly over time and across sampling sites, ranging from 0.2 to 100% (Kawata et al., 

2009; Nishizawa et al., 2009). The occurrence of herbicide-tolerance traits in feral canola 

populations in Japan is attributed to accidental loss and spillage of imported viable canola seed, 

of which approximately 90% is sourced from Canada, where up to 95% of the canola crop is 

comprised of transgenic HT varieties. Plants containing multiple herbicide-tolerant (a.k.a. 

stacked) traits (i.e., glyphosate- plus glufosinate-tolerant) have never been commercialized, but 

have been reported in feral canola populations (Aono et al., 2006; Kawata, 2010) as has the 

occurrence of herbicide-tolerance traits in interspecific (e.g., B. oleracea) and intergeneric (e.g., 

Sisymbrium altissimum; a Brassicaceae weed) hybrids with B. napus collected from roadsides in 

Japan (Kawata et al., 2009; Saji et al., 2005). However, it was noted that most hybrids with 

Sisymbrium sp. were sterile and did not produce seed (Kawata, 2010). Other surveys conducted 

in Japan have failed to detect the presence of herbicide-tolerance traits in seed collected from 

wild relatives of B. napus (e.g., B. rapa and B. juncea) sampled from ruderal habitats (Aono et 

al., 2006; Saji et al., 2005). At least in these two studies, the theoretical possibility of spilled 

feral transgenic herbicide-tolerant canola germinating, surviving, hybridizing with sexually 

compatible wild relatives and the hybrids surviving, reproducing and containing the transgene 

was below the level of detection.
Addendum to 73496 Canola petition Page 4 of 7

11-063-01p September 1, 2011

Table 11. Surveys to Monitor Transgene Presence in Feral Canola Populations

Country Surveyed area Period Transgene detection Sampled 

material

Belgium

Roadsides nearby and field 

margins of cropped fields in 

Wallonia

2007–2008 DNA analysis Leaf

Port areas (Antwerpen, Gent, 

Izegem and Kluisbergen)

Not 

specified DNA analysis Leaf

Canada*

Roadsides nearby and field 

margins of cropped fields in 

southern Manitoba (central 

Canada)

2004–2006

Herbicide screening, 

biochemical (protein) 

analysis

Seed, 

Leaf

Roadsides and railway lines in 

Saskatchewan and at the port 

of Vancouver

2005 Biochemical (protein) 

analysis Leaf

Japan

Port areas (Kashima, Chiba 

and Yokohama), roadsides and 

riverbanks in the Kanto district

2004

Herbicide screening, 

biochemical (protein) 

analysis, DNA 

analysis

Seed

Port areas, roadsides and 

riverbanks in western Japan 

(Shimizu, Yokkaichi, SakaiSenboku, Uno, Mizushima, 

Kita-Kyusyu and Hakata)

2005

Herbicide screening, 

biochemical (protein) 

analysis, DNA 

analysis

Seed

Port areas and roadsides in the 

area of Yokkaichi 2004–2007 Biochemical (protein) analysis Leaf

Roadside (Route 51) in eastern 

Japan 2005–2007

Biochemical (protein) 

analysis, DNA 

analysis

Leaf

USA*

Roadsides (interstate, state and 

country roads) in North Dakota 2010 Biochemical (protein) analysis Leaf

* Country where GE herbicide-tolerant canola is grown commercially

Source: adapted from Devos et al. (2011b)

Canola itself is not considered a noxious or invasive weed in the U.S., although it does 

demonstrate some of the common characteristics of weeds as described by Baker (1974); high 
Addendum to 73496 Canola petition Page 5 of 7

11-063-01p September 1, 2011

seed output under favorable conditions, continuous germination combined with long-lived seeds 

and self-compatibility, but not obligatorily self-pollinated or apomictic. However, canola has 

undergone many years of selective breeding and domestication and is a poor competitor with 

other species. Feral canola populations, even if genetically engineered, do not grow well outside 

of agricultural production.

Herbicide-tolerant canola is no more likely to form feral populations than unmodified canola, nor 

is it more likely to be more invasive or competitive or persistent in habitats where the target 

herbicide is not applied (Andersson and de Vicente, 2010; Warwick et al., 2009). Field studies 

in which both conventional and transgenic glufosinate-tolerant canola varieties were repeatedly 

introduced into 12 different habitats in the U.K. between 1990 and 1992, followed by monitoring 

over a period of up to ten years, failed to find any evidence that canola is invasive of natural 

habitats or that transgenic herbicide-tolerant canola is more invasive of, or more persistent in, 

disturbed habitats than unmodified canola (Crawley et al., 2001). As reported by Hall et al.

(2005), grower survey data from western Canada, where transgenic herbicide-tolerant canola has 

been grown extensively over many years, did not indicate any evidence of altered weediness or 

invasiveness potential.

The small size of canola seeds, their abundance and the tendency of canola seed pods to shatter 

means that canola is likely to appear as a volunteer in following crops and must be removed to 

preserve yield and crop quality. Control of volunteer canola plants can be accomplished with 

appropriate herbicides, taking into account the tolerance of the current crop and any tolerance of 

the previously planted canola variety. Failure to adequately control volunteers can lead to 

stacking of multiple herbicide-tolerance traits through cross pollinations, as has been observed in 

canola volunteers (Beckie et al., 2004; Hall et al., 2000) However, multiple herbicide-tolerant 

does not significantly alter plant fitness in the absence of herbicide pressure (Simard et al., 

2005).

Based on the lack of any alterations in survival and reproductive biology characteristics of 

currently commercialized transgenic herbicide-tolerant canola, Devos et al. (2011b) concluded 

that the likelihood of unintended environmental effects due to the establishment and spread of 

such plants will be no different than from conventional unmodified canola. The ability of 

transgenic herbicide-tolerant canola to successfully establish feral populations in disturbed 

habitats is limited by the availability of competition-free seed germination sites, just as for 

unmodified non-herbicide-tolerant canola.

The comparison of phenotypic characteristics between 73496 and non-transgenic canola 

demonstrated an absence of any biologically meaningful differences with respect to early 

population, vegetative growth, reproductive parameters, yield and ecological interactions. The 

73496 canola does not have an increased weediness potential and unconfined cultivation should 

not lead to increased weediness.

In unmanaged environments, herbicide-tolerant crops have no advantage over non-herbicidetolerant crops since no herbicide is applied. Based on this characterization, 73496 canola is no 

more likely to establish feral populations than either existing transgenic herbicide-tolerant canola 
Addendum to 73496 Canola petition Page 6 of 7

11-063-01p September 1, 2011

varieties or unmodified non-herbicide-tolerant canola, and these populations would not be more 

likely to be persistent or invasive.

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