White Paper

on

RNAi Technology as a Pesticide: Problem Formulation for Human Health and Ecological 

Risk Assessment

Submitted to the FIFRA Science Advisory Panel

For Review and Comment

October 29, 2013

U.S. Environmental Protection Agency

Office of Chemical Safety and Pollution Prevention

Office of Pesticide Programs

Biopesticides and Pollution Prevention Division

[September 30, 2013]
RNA-Interference FIFRA Scientific Advisory Panel White Paper

In the past several years, a number of pharmaceutical and agricultural products based on RNA 

interference have been developed. Most of these products are pharmaceuticals aimed at turning 

off aberrant protein expression (e.g., cancer genes or macular degeneration), but a few

agricultural products utilizing RNA interference have also been developed for pest control. The 

potential for increasing use of RNA interference to control pests has lead EPA to conclude the 

Agency should convene a Scientific Advisory Panel (SAP) to consider aspects of risk assessment 

the Agency uses to determine whether an RNA interference based PIP can meet the 

environmental safety standard of the Federal Insecticide Fungicide and Rodenticide Act 

(FIFRA). 

EPA is not requesting of the SAP a review of any of the Agency’s past or current decisions 

related to RNA interference based products, but would appreciate a consultative discussion on 

issues that might arise as the technology develops and finds wider application in the future. 

Given the likelihood for development of both plant expressed RNA interference products for pest 

control (i.e., RNAi in Plant-Incorporated Protectants or PIPs) and other types of dsRNA pest 

control products (i.e., non-PIP dsRNA end use products, e.g., the dsRNA the Agency views as a 

biochemical), EPA addresses both types of products in this White Paper, and requests discussion

on both types from the SAP.

I. Background: Agricultural Products Based on RNA Interference with an Emphasis on 

Those Reviewed by EPA

The FlavrSavrTM tomato, one of the first plant products of modern biotechnology to be approved 

for human consumption in 1992 by the Food and Drug Administration, , was developed using 

what was then termed “anti-sense” technology. The FlavrSavrTM tomato was engineered using 

the antisense technology to turn off expression of a polygalacturonase enzyme involved in the 

softening of the fruit during ripening. In 1998, a potato engineered to express the gene for potato 

leaf roll virus replicase was registered as a PIP by EPA (EPA, 1998a). This product differed

from the products previously examined by EPA in that the company could not detect any 

replicase protein being expressed, yet its stable phenotype1

was resistant to viral infection. The 

resistance traits of these plant transformations have subsequently been theorized as being due to 

the process of RNA interference. 

More recently, EPA registered another PIP expressing a transgene which delivers viral resistance 

in European plum to the plum pox virus via an RNAi pathway; this PIP expresses a coat protein 

1 EPA was able to make a dietary safety finding for this virus resistant PIP based on the fact that 

virus infected produce is known to long be part of the food supply without any untoward effect 

on human and domestic animal health.

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gene from the plum pox virus (EPA, 2010a). This PIP does not produce detectable quantities of 

coat protein as the coat protein transgene functions to suppress infection by the plum pox virus. 

In 2013, the EPA approved an experimental use permit (No. 524-EUP-104) for field evaluation 

of corn engineered to express a transgene construct which includes the Snf7 gene which 

suppresses mRNA encoding vacuolar ATPase subunits A and E (Bachman et al., 2013). This 

PIP targets the Western corn rootworm, and published data indicate the specificity of the Snf7

gene to the target pest ATPase mRNAs with no activity noted toward the other insect species 

evaluated. 

II. RNA Interference – Overview

Interference or suppression of gene expression by naturally occurring double stranded 

ribonucleic acid-based mechanisms was largely unknown or misunderstood until the work of 

Andrew Fire and Craig Mello (Fire et al., 1998) which earned them the Nobel Prize in 

Physiology or Medicine in 2006. Previous efforts to modify plants, insects, mammals and other 

organisms with transgenes or antisense constructs producing single stranded RNA (ssRNA) were 

well documented (Izant and Weintraub, 1984; Fire et al., 1991; Lee et al., 1993; Guo and 

Kemphues, 1995; Matzke and Matzke, 1995; Goodwin et al., 1996; Akhmanova et al. 1997; 

Taylor, 1997), however, the discovery that double stranded RNA (dsRNA) was 10 times or more 

potent in its effect on gene expression than ssRNA unleashed a wealth of research into the 

biochemical basis for this mechanism known as RNA interference (RNAi) (Fire et al, 1998). 

Post-Translational Gene Silencing (PTGS) or co-suppression was noted in many instances in the 

late 1980s and 1990s wherein transgenic sequences introduced into plant genomes often resulted 

in transient expression followed by silencing and return to the untransformed phenotype (de 

Carvalho et al., 1992; Finnegan et al., 1994; Flavell, 1994). Plant resistance to RNA viruses had 

been demonstrated to function well when DNA sequences matching the sequence of coat protein

or other viral genes were introduced into plants as transgenes; however the mechanisms for both 

observations remained nebulous, although many theories were proposed. 

Working with Caenorhabditis elegans, a common saprobic nematode, the laboratories of 

Andrew Fire and Craig Mello ascertained that injected dsRNA sequences matching known gene 

sequences in the nematode genome was quite effective at gene silencing (Fire et al., 1998; 

Grishok and Mello, 2002). Injection of dsRNA sequence matching a mex-3 gene of C. elegans

resulted in an absence of mex-3 mRNA and an alteration in phenotype (i.e., embryonic arrest) in 

contrast to ssRNA injections which yielded little change in corresponding mRNA populations 

and > 90% embryo production. 

Evidence for naturally occurring gene silencing, and that resulting from introduction of foreign 

DNA or RNA sequences, has been noted in numerous prokaryotic and eukaryotic organisms to 

date (Waterhouse et al., 1998; Makarova et al., 2006; Pridgeon et al., 2008; Auer and Frederick, 

2009; Huvenne and Smagghe, 2010; Younger and Corey, 2011; Adamo et al., 2012). As 

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research continues utilizing RNA interference (RNAi) for elucidation of biochemical pathways, 

genomic structure analysis and for practical applications like pest or disease resistance in plants, 

insects and humans, it is clear that differences exist at the molecular level regarding the details of 

the RNAi pathway to gene silencing between groups of organisms and sometimes even within 

species of a single genus. 

A. General Features of the RNA Interference Pathway

While differences exist among organisms in the biochemical details of the RNAi pathway, some 

general features of the mechanism appear to be conserved. The various forms of dsRNA clearly 

show differences in structure and in initial production / processing stages; however, they also 

share a common mode of action. All of these dsRNAs are processed by Dicer or a Dicer-like 

nuclease, to yield small-interfering RNAs (siRNAs) which then attach to an Argonaute (AGO) 

family protein preferentially binding to one (guide strand) of the two strands of RNA in the 20 to 

30 nucleotide (nt) molecule; the other RNA strand or passenger strand is typically degraded 

following release from AGO in the RNA Induced Silencing Complex (RISC). AGO is an RNAse 

H type enzyme, an endonuclease, which degrades mRNAs with complementarity to the bound 

miRNA or siRNA sequence. This activity, which utilizes the guide strand to target and bind 

complementary mRNA sequences for subsequent degradation is referred to as Slicer activity. 

Long dsRNA molecules are thought to be critical to triggering RNA interference. The long

strands of dsRNA that trigger RNA interference can originate outside the cell or be internally 

produced. Small interfering RNA is the product of Dicer activity on longer molecules of dsRNA. 

The siRNA molecules are also double stranded from 21-24 nucleotides (nt) in size with a 2 nt 

overhang at the 3’end. These were first described in studies of transgene-induced and viralinduced silencing in plants (Hamilton and Baulcombe, 1999; De Paoli et al. 2009). siRNAs may 

be processed from long dsRNAs and short hairpin RNAs, as well as being exogenously supplied 

by injection into a cell or via transformation or transfection. In addition to their role in RNAi 

through PTGS, siRNAs are known to effect chromatin modifications (e.g., cytosine or histone 

methylation, histone acetylation).

There are also other forms of RNA molecules triggering RNAi that can be generated internally 

from the host chromosome such as the micro RNA (miRNA), PIWI-interacting RNA (piRNA) 

and trans-acting RNA (tasiRNA). While all these RNA types are stated as being separate 

entities, they are similar in structure when involved in the RNA interference pathway. All these 

RNA forms are processed by cellular systems that result in short 21-31 nt single stranded RNA 

guide molecules loaded into the RNA induced silencing complex (RISC) (e.g., Argonaute or its 

homologues with perhaps other associated enzymatic activities; Pratt and MacRae 2009). The 

RISC enzymatic complex is guided by the 21-31 nt guide sequence loaded into Argonaute for 

specific recognition, binding and activity toward a target mRNA molecule. In the instances 

involving siRNA and miRNA, AGO is a part of RISC, which achieves gene silencing by 

degradation of the corresponding mRNA or by transcriptional repression (Wilson and Doudna, 

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2013). The outcome of this binding can be either direct cleavage of the target mRNA by RISC or 

repression of the targeted mRNA translation. The piRNA and tasiRNA have been associated 

with chromatin restructuring. The overall effect of RNAi is suppression of the targeted gene’s 

phenotype.

siRNAs rely on a high degree of specificity or complementarity based upon sequence alignment 

with the target mRNA. miRNAs, in contrast, do not always require the same degree of base 

sequence complementarity in order to effect gene silencing through either mRNA degradation or 

chromatin modification. 

dsRNA appears to be a key element in RNAi but there are several sources of the dsRNA. 

dsRNA molecules may be produced within the nucleus of a cell in the form of pri-miRNA 

(primary micro RNA) which are produced by RNA Polymerase II activity on a host gene and

are approximately 1000 nt long. These long dsRNAs are processed by a ribonuclease III 

(RNAse) enzyme known as Drosha and a dsRNA binding protein known as Pasha (in 

invertebrates) or DGRC8 (DiGeorge Syndrome Critical Region 8; in mammals), collectively 

referred to as the microprocessor complex, to form pre-miRNA molecules. These pre-miRNA

(precursor miRNA) molecules, approximately 70 nt in length and containing stem and loop 

structures, are exported to the cytoplasm via the nuclear pore complex and bind with a RNAse III 

enzyme called Dicer (or Dicer-Like 1 (DCL1) in plants; Dcr1 and Dcr2 in insects). This enzyme 

cleaves the RNA sequence into short sequences, miRNAs, typically 21-25 nt in length. This 

binding to Dicer also initiates association with the larger RISC complex where binding of 

corresponding mRNAs will ultimately take place. Interestingly in plants, the DCL1 activity 

occurs in the nucleus and not the cytoplasm. For animal miRNAs, (vertebrates and some 

invertebrates) the two reactions are split spatially, one in the nucleus and one in the cytoplasm. 

To aid in accurate processing of plant miRNAs and RNA strand selection, an RNA binding 

protein HYPONASTIC LEAVES 1 (HYL1) and the zinc finger protein SERRATE (SE) function 

coordinately with DCL enzymes (Manavella et al., 2012a).

piRNAs (PIWI-interacting RNAs) are short ssRNAs (26-31 nt) present in vertebrates and some 

invertebrates, found in both the nucleus and cytoplasm whose biogenesis is unclear as they do 

not require Dicer to produce an effect. They are known to cause gene silencing, often targeting 

retrotransposon sequences, and also have a role in epigenetic phenomena which may be 

maternally heritable. piRNA are especially present in germ-line cells and are known to influence 

spermatogenesis. 

Within the group of small dsRNAs known as siRNAs, transacting siRNAs (tasiRNAs) are 

endogenous RNAs which down regulate genes at loci different from which they arose (Peragine 

et al., 2004; Vasquez et al., 2004). TAS genes, loci from which tasiRNAs are derived, are known 

from work in Arabidopsis thaliana, Oryza sativa, other higher plants and moss, but have not yet 

been identified in animals or fungi. While they have some similarities with miRNAs and other 

siRNAs in that they post-transcriptionally silence gene expression, the biogenesis of tasiRNAs 

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differs in the requirement for RNA-dependent RNA polymerases (RDRP) and SGS3 (suppressor 

of gene silencing), known to be present in plants in multiple forms. One or more forms of DCL 

enzymes are also required for proper processing to the final 21 nt length, as are components 

typical of the miRNA pathway (e.g., AGO1, HEN1, HYL1). tasiRNAs are derived from longer 

dsRNAs as compared to miRNAs.

TasiRNAs are incorporated into the RISC complex and thereby direct the cleavage of target 

mRNAs, albeit with less fidelity relative to sequence alignments, to degrade mRNA. This 

reduced sequence specificity in tasiRNAs is shared by some miRNAs as well. It has been 

demonstrated in plants that miRNA directed cleavage of target mRNA sequences can lead to the 

production of secondary siRNAs which can lead to the degradation of non-targeted mRNA 

through the formation of transitive RNAs (Himber et al., 2003; Manavella et al., 2012b). 

Transitivity can be triggered by the specific AGO involved in the RISC complex as well as the 

22 nt length of the guide strand retained in the effector complex. Additionally, asymmetrical 

bulges in the miRNA:miRNA* duplex resulting from a lack of complete complementarity can 

trigger transitivity. Why some miRNAs trigger transitivity and some do not is not well 

understood at this time. A similar transitive effect has been seen in C. elegans following 

amplification of new dsRNA and a resulting non-autonomous effect (Alder et. al., 2003; 

Roignant et al., 2003). That is, the silencing of a gene targeted in one cell can lead to the 

silencing of a second gene in a distinct cell type. This phenomenon has not been noted in 

Drosophila and its absence attributed to the lack of RdRP activity in this insect (Roignant et al., 

2003). 

Small RNAs which act upon highly related sequences (mRNAs) are referred to as auto-silencing 

RNAs whereas small RNAs which act upon sequences derived from largely unrelated loci with 

minimal sequence similarity are referred to as hetero-silencing RNAs (Vasquez et al., 2004). The 

latter are termed trans-acting in that they target mRNAs from genes to which they bear little 

resemblance and the siRNAs which target closely related and highly homologous sequences are 

termed cis-acting RNAs.

In addition to the modulation of gene expression via mRNA degradation (e.g., PTGS), as 

directed by some miRNAs and siRNAs, heterochromatic siRNAs modify histone proteins and 

DNA (cytosine) associated with heterochromatin in insects and fungi (Reinhart and Bartel, 

2002). By altering chromatin rather than mRNA, these siRNAs influence gene expression at the 

transcriptional level. These siRNAs derive from endogenous sequences through dsRNA 

precursors and are processed by Dicer, then bind to a RNA-Induced Transcriptional Silencing 

(RITS) complex, at least in Drosophila and Schizosaccharomyces pombe (Fagegaltier et al., 

2009). RITS complexes contain Argonaute, a chromodomain protein Chp1, and Tas3, and they 

interact with a RNA-directed RNA polymerase, a polyA polymerase (Cid12) and a helicase 

(Hrr1), termed RDRC, necessary for the formation of dsRNA. Both RITS and RDRC interact 

with noncoding RNA transcripts derived from pericentromeric areas of the chromosome to 

trigger the RNA interference pathway and assembly of heterochromatin. A methyltransferase 

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(Clr4) aids in the association of RITS and RDRC with chromatin and is in part responsible for 

the formation of siRNA. Methylation of histone protein 3 is mediated by this methyltransferase 

in Schizosaccharomyces pombe, the fission yeast; methylation of histone proteins and cytosine 

bases can provide transgenerational alteration of phenotype through their impacts on gene 

expression of select chromosomal regions.

Ago1, RDRC and Dicer are all required in the formation of siRNAs from specific noncoding 

RNA transcripts (Halic and Moazed, 2010). In the absence of Dicer, transcriptomal degradation 

products termed primal RNAs interact with Ago1 to drive the initial formation and amplification 

of siRNA in the absence of heterochromatin (Conte and Mello 2010). Interestingly, it appears 

that heterochromatin formation in humans and other mammals is independent of the RNAi 

pathway.

B. Post-Transcriptional Repression Through Deadenylation 

The RNAi pathway in plants, fungi and animals also includes post-transcriptional repression of 

mRNA translation through decapping and deadenylation pathways which serve to decrease the 

stability of mRNAs and to trigger their ultimate degradation. Three different types of 

deadenylase complexes (i.e., PARNS, CCR4-Not, PAN) are known to deadenylate mRNA 

following shortening of the 3’ poly-A tail. As the tail is shortened, the mRNA typically 

undergoes decay by resident exoribonucleases, present in the cell cytoplasm in plants (Abassi et 

al., 2013). The removal of the 5’ 7-methylguanylate cap (decapping) may or may not be linked to 

the 3’-5’ degradation of mRNAs, however, it does preceed the 5’-3’ degradation of mRNA 

sequences by exonucleases such as XRN1. In mammals, miRNAs are known to bind to the 5’ 

untranslated region (UTR) of the mRNA which is often followed by deadenylation and mRNA 

decay (Beilharz et al., 2009). miRNAs are also known to effect translational repression of 

mRNAs in Drosophila and zebrafish (Danio rerio) through deadenylation and subsequent 

destabilization of mRNA. 

C. RNAi and Cellular Maintenance

RNAi pathways can be seen as a normal part of cell maintenance and as a response to stress 

events (e.g., viral infection, abiotic stress) which seek to maintain homeostasis. Some have 

characterized RNAi pathways as a surveillance system to prevent translation of aberrant mRNAs 

and other nucleic acid sequences (e.g., viruses, transposons) (Zamore et al., 2000). Although the 

details of the RNAi pathways and their outcomes may differ among organisms, what is clear is 

that the influence of small RNAs on growth, development, defense and even transient heritability 

of traits is substantial. With that in mind, the potential for modification or manipulation of these 

pathways for pest management, agronomic production and essentially any other desired 

phenotypic alteration is great and will lead to a variety of approaches to delivery of small RNAs.

siRNAs are often delivered to plant or animal cells from exogenous sources, although, as 

described above, they can also arise from the host genome (i.e., miRNA, tasiRNA, 

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heterochromatic RNA). While most miRNAs arise from endogenous DNA sequences, efforts are 

underway to manipulate these sequences to effect the targeting of select mRNAs. Early 

experiments which helped divulge the existence of the RNAi pathway relied upon soaking in 

dsRNAs or ingestion of these dsRNAs by C. elegans (Fire et al., 1998). Injection of small RNAs 

into the gonads of C. elegans or tissues of other organisms has also been a well used delivery 

procedure for research purposes, however, not as a practical approach for agricultural uses. 

Plant transformation, that is the introduction of novel nucleic acid sequences into the plant 

genome, is an obvious delivery system for expression of novel traits in plants, including dsRNAs 

(Bachman et al., 2013). However, delivery of dsRNAs to plants, insects or other organisms via 

spraying with dsRNA (Wang et al., 2011) or being carried in a microbe (Zhu et al., 2011) are 

also feasible approaches depending on the target organism and ability of that organisms to take 

up the RNA directly. Oral ingestion through feeding is a common approach to test the RNAi 

response in insects (Turner et al., 2006). In C. elegans, Sid-1 (Systemic RNAi Defective) has 

been shown to be critical to dsRNA uptake in some instances and capable of effecting systemic 

spread of the RNAi signal (Gu and Knipple, 2013). Receptor mediated endocytosis has also been 

demonstrated in these two organisms for uptake of dsRNA from the gut lumen. 

The fate and impact of small dsRNAs on organisms has been divided into three principle 

groupings: cell autonomous, environmental and systemic (Gu and Knipple, 2013). Cell 

autonomous RNAi refers to instances wherein the effect of the dsRNA is in the cells where the 

dsRNA was produced or exogenously supplied and is therefore limited spatially. Environmental 

RNAi impacts result from exogenous uptake from the immediate environment, such as an insect 

gut or hemocoel, or even from incidental contact in the soil. If the silencing effect of the dsRNA 

continues to spread among neighboring cells, even cells in which the target RNA does not exist, 

this is referred to as systemic RNAi. Environmental and systemic RNAi are termed non-cell 

autonomous since the dsRNA does not have to be produced at the site (cell) of action. 

RDRP is present and active in C. elegans, notably amplifying the level of siRNAs and aiding in 

the systemic RNAi response (Sijen at al., 2001). It is known, however, that siRNA amplification 

and even core RNAi components are not necessary for systemic RNAi induction (Tomoyasu et 

al., 2008). Among insect species, much of the RNAi machinery is largely conserved, although 

responses to dsRNA, including systemic spread are highly variable (Aronstein et al., 2011; 

Terenius et al., 2011). RDRP is not known from insects although one cannot rule out the 

presence of alternative enzyme activities with overall little direct homology to RDRP as known 

from other organisms.

As discussed further in the sections to follow, the EPA is formulating a risk assessment approach 

for products using RNAi-based pathways to perform its duties under the Federal Insecticide 

Fungicide and Rodenticide Act (FIFRA), the Food Quality Protection Act (FQPA) and the 

Federal Food Drug and Cosmetic Act (FFDCA) relative to the safe use of a pesticide and any 

8
impacts upon man and the environment. Uncertainties clearly exist with respect to a complete 

understanding of all current and future applications of this technology, however, the Agency’s 

previous experience with RNAi-based PIPs and attention to ongoing developments in research 

on small RNAs provides a basis for such assessment. This FIFRA Scientific Advisory Panel will 

examine potential impacts upon humans from consumption of RNAi-based PIP and non-PIP 

products, interactions of dsRNA with non-target organisms, and the environmental fate of 

dsRNA when introduced as part of a PIP or as a direct spray application of dsRNA for pest 

management. The questions proposed to this panel will help to focus the discussion on the 

Agency’s needs relative to data gaps which may exist. 

III. Mammalian Safety Assessment

The current risk assessment for mammalian safety for PIP and biochemical products utilizes data 

and information determined on a case by case basis. The product characterization information 

accurately describes the pesticidal product, its original source, its mode of action, how it was 

introduced and how it can be detected. For biochemical pesticides, there is an additional 

discussion of the manufacturing method, including consideration of any materials used during 

manufacture or added to formulate the final product. 

The hazard analysis of PIPs depends on information about the specific amino acid sequence and 

comparative analyses using computer algorithms to discover any significant sequence similarities 

to known toxins or allergens. There is also basic chemical information about the stability of the 

protein to heat and digestive enzyme activity. Finally, there is toxicity information from studies 

performed with a purified protein or from information in the peer-reviewed scientific literature. 

The studies for mammalian safety determinations of PIP proteins utilize the microbial guidelines 

(40 CFR 158.2140) and typically consist of a single acute toxicity study and the in vitro and 

bioinformatic analyses. For biochemical pesticides, the hazard analysis is based on the tiered 

testing scheme described in the guidelines (40CFR 158.2050) which considers the results of 

toxicity studies along with information from the peer-reviewed scientific literature to make a 

safety finding. 

Safety determinations for non-PIP dsRNA products may need to rely on selected guideline 

studies under 40CFR 158.2050 due to likely stability differences between PIP and non-PIP 

dsRNAs. It is anticipated that non-PIP dsRNAs that are exogenously applied to target pests or 

targeted use sites, will be formulated into end-use products (EPs) in a manner that will increase 

their stability to abiotic and biotic degradation in the environment (see a more detailed discussion 

in Section IV.B. of this document), to provide sufficient residence time in/on the treated articles 

for the dsRNA to accomplish the desired pesticidal activity. The presence of “stabilized” 

dsRNA in/on treated articles increases the potential opportunities for topical, inhalation, and 

9
dietary exposure that can be assessed with the traditional Tier I hazard testing for biochemical 

pesticides. These studies are listed below:

• Acute oral toxicity

• Acute dermal toxicity

• Acute inhalation toxicity

• Acute eye irritation

• Acute dermal irritation

Depending upon the residence time of “stabilized” dsRNA following application, additional 

longer term studies may be needed, such as:

• 90-oral toxicity

• 90-day dermal toxicity

• 90-day inhalation toxicity

Finally, an exposure assessment is performed on the product based on its intended use, if the 

results of the hazard analysis warrant that consideration.

A general approach to the risk assessment of mammalian health effects for dsRNA molecules 

used for pest control resembles the process described above for protein PIPs and biochemicals

but can be further developed based on two considerations. These considerations are directed 

towards factors that might mitigate hazards and limit possible exposure. The first is the design 

of dsRNA sequences to control pests. Appropriate sequence selection to trigger pest control may 

lessen the potential for affecting non-targeted species. A thorough bioinformatics search and 

comparison should be performed to insure specificity of the pest control sequence; i.e., an 

appropriate sequence should display no significant similarity to other non-targeted gene 

sequences, especially those found in mammalian species. The second consideration is the low 

likelihood that any significant mammalian exposure could result from dietary or other anticipated 

exposures (e.g., dermal or inhalation) once the digestive degradation and environmental fate of 

dsRNA is examined. These suppositions need to be made explicit and examined for supporting 

data. Information supporting these suppositions include: (1) an understanding of the molecular 

basis of the RNA interference phenomenon as it relates to the safety assessment; (2) results of 

human therapeutic research on the use of RNAi technology to treat human disease, and (3) 

research indicating the level of sequence specificity and/or amount of dsRNA necessary to 

trigger successful RNA interference.

A. A Brief Overview of the Molecular Basis of the RNA Interference (RNAi) 

Phenomena as it Informs the Mammalian Safety Assessment.

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As noted in the previous section of this White Paper, there are three forms of dsRNA molecules 

that are generally acknowledged to be involved in RNA interference in mammals: micro RNA 

(miRNA), PIWI interacting RNA (piRNA) and small interfering RNA (siRNA). miRNA is 

transcribed from the genome as a pri-miRNA then processed and exported from the nucleus as 

pre-miRNA with a distinctive small hairpin or stem-loop structure. The piRNA is involved in 

silencing transposons during germ cell formation. The siRNA is the product of Dicer activity on 

longer molecules of dsRNA. These longer strands of dsRNA can originate outside the cell or be 

internally produced. The siRNA molecules are also double stranded from 21-24 nucleotides (nt) 

in size with a 2 nt overhang at the 3’end. While all three of these RNA types are stated as being 

separate entities, they are similar in structure when involved in the RNA interference pathway. 

All these RNA forms are processed by cellular systems that result in short 21-31 nt single 

stranded RNA guide molecules loaded into the RNA induced silencing complex (RISC) (e.g., 

Argonaute or its homologues with perhaps other associated enzymatic activities) . The RISC 

enzymatic complex is guided by the 21-31nt sequence loaded into Argonaute for specific 

recognition, binding and activity toward a target mRNA molecule. The outcome of this binding 

can be either direct cleavage of the target mRNA by a slicer activity in the RISC complex or 

repression of the targeted mRNA translation. The overall effect of RNAi is suppression of the 

targeted gene’s phenotype.

Differences between the miRNA, piRNA and siRNA are critical for their expected function. 

miRNAs originate in the nucleus and have the secondary structure of a hairpin or stem loop with 

the dsRNA sections that can be processed into 21-24 nt siRNAs. These smaller siRNAs can 

then be loaded into the RISC complex and have mostly host developmental functions of up or 

down regulation of the host genome. piRNA also originates in the nucleus, does not have such a 

complex secondary structure, may not be double stranded prior to loading into the RISC complex 

as 26-31 nt species and functions mostly to inactivate transposons in germ line cells, especially 

those associated with mammalian spermatogenesis. siRNA is thought to be the product of Dicer

activity on longer dsRNA molecules, is generally 21-24 nt when functional in the RISC complex 

and is believed to have higher sequence complementarity to trigger mRNA cleavage. Both 

miRNA and piRNA require less sequence complementarity than siRNA and have been more 

frequently involved in translational pausing in mammals and higher organisms rather than direct 

mRNA cleavage.

The RNA interference phenomenon is found in most forms of life but there are significant 

differences in some of the components depending on the taxonomic kingdom or group. One 

significant difference has been the presence of an RNA dependent RNA polyermase (RdRP) in 

plants and nematodes but not in vertebrates. RdRP is responsible for increasing the number of 

dsRNA molecules present and for the systemic spread of RNAi in plants. Systemic spread has 

been associated with viral disease resistance and other systemic signaling effects in plants. The 

lack of systemic spread in vertebrates has made delivery of therapeutic agents for RNA 

interference more challenging.

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B. Therapeutic Research in Humans

In mammals, and humans in particular, there has been a great deal of research to describe the 

RNAi system which has resulted in information that is cogent to dietary risk assessment. There 

has been extensive research using RNAi for reverse genetics to examine the effects of trait 

silencing. The most relevant use of RNAi applicable for the mammalian health risk assessment 

has been the development of pharmaceutical drugs to treat diseases by silencing specific cellular 

functions. The dsRNA based pharmaceuticals have generally been shown to be effective in cell 

culture studies but delivery to the target organ by the oral, dermal or inhalation route has been 

difficult (Castanotto and Rossi, 2009). Most dsRNA therapies based on native forms of dsRNA 

have not shown efficacy by the oral route of administration without some protection from 

exposure to stomach acids and RNases (Petrick et al., 2013). For dermal and inhalation routes of 

drug delivery, some form of conjugation or encapsulation is necessary to facilitate 

transmembrane movement (Castanotto and Rossi, 2009). Some RNAi interventions have also 

shown off-target effects on the treated animal where other genome sequences were silenced 

along with the target sequence and immunostimulatory effects, particularly on the innate immune 

system have been noted (Whitehead et al., 2011; Meng et al., 2013). It is important to note that 

since there is no new protein being expressed, the use of RNA interference technology, should

not present any allergenicity issues. The use of RNAi as PIPs presents a lower exposure scenario 

by all routes for mammals compared to the stabilized and sprayable formulations of dsRNA (see 

section V. on non-PIP dsRNA end-products)

C. Open Questions on Sequence Specificity 

The implication that sequence specificity of the dsRNA molecule drives the recognition and 

subsequent degradation of the target mRNA should allow a robust bioinformatic analysis of the 

proposed target gene and the dsRNA product. EPA has not yet established what form this 

bioinformatic analysis should take and what level of similarity would trigger concern for offtarget effects. There are already reports that rice, corn and other crop plant species have siRNA 

species present that have high homology to human gene sequences (Ivashuta et al., 2009). These 

represent similarities in the 21-24 nt range which would not be unexpected given simple 

probability analysis for such short sequences. The significance of the random short homologies 

versus longer contiguous homologies has not been examined. However, if significant 

homologies of the overall dsRNA product are found with mammalian genes or regulatory 

regions, it would be prudent to examine the dsRNA molecules potential effect more closely. The 

specificity of the Dicer enzyme in the generation of siRNA species is not clear so that the need 

for a 21nt stepwise or other modified bioinformatic analysis to predict off-target effects is still 

unsubstantiated. The presence of a 3’ overhang of one to three nucleotides is a dominant force 

for the specificity of Dicer activity but longer overhangs or blunt ends will also generate the 

siRNA species (Vermulean, 2005).

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Indications are that the majority, if not all, orally administered dsRNA is degraded by the acid 

environment of the stomach and the action of pancreatic RNases (Rehman et al., 2011; 

Sorrentino et al., 2003; Hoerter et al., 2011). This degradation should lessen the probability that 

any dsRNA PIP could survive gut passage and be taken up at a concentration that would trigger 

RNAi. Cell culture studies indicate that at least 100 copies of siRNA molecules need to reach a 

targeted cell site to induce RNAi in mammalian cells (Brown et al., 2007). If the need for high 

copy number delivery to target cells is accurate, there may also be levels of RNA molecules with 

potential similarity to mammalian genes that could be safely consumed without triggering RNA 

interference. 

However, there are still questions of the stability of individual dsRNA molecules and if 

chemically modified forms or RNA molecules with significant secondary structure such as 

hairpins or super coils, could survive acid and enzymatic degradation leading to greater 

exposure. There are reports in the literature that miRNA molecules from plants consumed in the 

diet have been absorbed and shown to induce RNAi in humans (Zhang et al., 2012). There have 

also been subsequent reports that tend to refute the findings of significant uptake of miRNA from 

the diet (Witwer, 2013; Snow, 2013). There are also several reports on the lack of successful 

development of RNA based therapeutic drugs for oral delivery which also question the dietary 

exposure and uptake of RNA molecules (Petrick et al., 2013; Castanotto and Rossi, 2009).

IV. Environmental Risk Assessment

Double-stranded RNA may be utilized as a pesticide that is expressed as a plant incorporated 

protectant (PIP), or it may be applied to the plant or the environment in the form of a foliar 

spray, post-directed spray or drench, seed treatment, or granule/powder. While exposure 

scenarios may differ between dsRNA PIPs and exogenously applied dsRNAs, EPA would have 

similar considerations for environmental fate of dsRNAs and nontarget effects under the 

different exposure scenarios. The sections below discuss the unique issues relevant to exposure 

of nontarget organisms to dsRNA PIPs and exogenously applied dsRNAs, as well as the 

uncertainties related to the current approaches to risk assessment.

A. Environmental Considerations for PIP-Expressed dsRNA

Over the past two decades, ecological risk assessments have been performed for numerous PIPs, 

most of which express proteins derived from the bacterium Bacillus thuringiensis (Bt, e.g., see 

EPA, 2005; 2010b; 2010c). The current approach to ecological risk assessment for Bt PIPs 

provides a framework for risk assessment of dsRNA PIPs. However, as this section will 

describe, not all aspects of the fate of dsRNA in the environment and potential effects on 

nontarget organisms are necessarily understood. As a result, dsRNA PIPs may present unique 

challenges for ecological risk assessment that have not yet been encountered for other PIPs. 

13
This section discusses issues and uncertainties related to the two major components of ecological 

risk assessment as these relate to dsRNA PIPs, which are the estimation of environmental 

exposure and determination of effects resulting from exposure. Details of the approach generally

used by EPA to assess ecological risk for other types of PIPs (e.g., see EPA, 2005; 2010b; 

2010c) are presented herein only where potential differences of approach may be required for the 

risk assessment for dsRNA PIPs. Some aspects of ecological risk assessment commonly 

considered for PIPs, such as potential for gene flow through outcrossing, potential for 

development of invasiveness, and horizontal gene transfer, are not discussed because EPA 

assumes that the current approach that has been used for Bt PIPs would also apply to dsRNA. 

The discussion in this section is focused on to assessing risks dsRNA PIPs may present to 

nontarget organisms.

1. Nontarget Exposure

Important considerations in any environmental risk assessment are the stability of the pesticidal 

substance, in this case the dsRNA, and the potential routes through which the pesticidal 

substance may be distributed to the environment. As this section of this issue paper examines 

dsRNA that is tightly integrated into the plant, i.e., is a PIP, the analysis will focus on the 

expression and stability of dsRNA in the plant and how dsRNA integrated into plant tissue may 

persist or move through the environment, thus creating the conditions of potential exposure to 

nontarget organisms.

The degree of nontarget exposure depends on the distribution and fate of the dsRNA PIP within 

the environment, as well as the potential routes of exposure, exposure duration, and potential for 

uptake. These factors are discussed here, as well as qualities unique to dsRNA PIPs that may 

introduce additional uncertainty into exposure estimates.

a. dsRNA Distribution and Fate in the Environment

 

Factors that affect the distribution of dsRNA in the environment and the potential for exposure to 

nontarget organisms include physical movement of the PIP crop plant tissue, persistence and 

physical movement of dsRNA released from the plant, either while the plant is living or 

following breakdown of plant tissue. EPA assumes that the dsRNA expressed in a PIP crop plant 

would be present wherever any of that crop plant tissue exists or is moved unless sufficient data 

are presented to show otherwise. On the crop field, living plant tissue will be present both above 

and below the soil surface where the crop plants are grown. During the growing season, plant 

tissue may be moved within the field through pollen release or the breakage and physical 

movement of other plant tissues (e.g., leaves) from the plants. Ultimately, these plant tissues are 

expected to reach the soil surface, though pollen may fall onto plant surfaces first. Once the plant 

is harvested, crop debris could be deposited on the ground surface or tilled into the soil. Root 

14
tissue is expected to remain in the soil until it breaks down or is harvested, except in the case of 

perennial plant for which living root tissue would persist. 

It is unclear at this point whether a dsRNA PIP also would be incidentally present in root 

exudates, guttation droplets, or nectar, providing additional on-field sources of nontarget 

exposure. This may be affected by the characteristics of the dsRNA or the plant. Most PIPs target 

insect pests that chew plant tissue, and exposure considerations thus have been focused on 

nontarget organisms that eat solid plant tissues. However, some current work is focused on using 

dsRNA expressed in planta to control plant hoppers and other sucking insects (Pitino et al., 

2011; Zha et al., 2011), and Mlotshwa et al. (2002) indicate that systemic gene silencing may 

occur as a result of movement of the silencing signal in phloem, although the exact mechanism is 

not confirmed. These studies suggest that these sources of exposure may also need to be 

considered in a risk assessment. 

Off-site movement of plant tissue is also possible. Depending on whether the dsRNA is 

expressed in the pollen, significant consideration may be given to the role pollen may play in offfield nontarget exposure. Some pollen is expected to move off site, and the amount and the 

distance moved will depend on the characteristics of the pollen (e.g., morphology, weight) and 

the mechanism relied upon for pollination (e.g., wind, pollinators, self-pollination). Animalpollinated crops would be expected to play a smaller role in off-field nontarget exposure, and 

concerns in these cases would be more focused on the exposure of the pollinators themselves. 

Other plant tissues are also expected to move off site, and an assessment would consider whether 

these tissues could contribute significantly to nontarget exposure, as well. In some crops, postharvest plant debris may be left on the field and in the immediate field margins. Debris may be 

moved further away from the field in terrestrial environments by wind, water, or another 

mechanism (e.g., physical movement by man), and may also enter aquatic systems where it could 

remain near the site of entry or be moved by water currents. Additionally, harvest of some crops 

can create dust out of dried plant material, which may move offsite. For determining the extent 

and degree of off-field nontarget exposure related to plant tissue movement, EPA would consider 

the expression and stability of the dsRNA PIP within plant tissue, and may also rely on data on 

pollen movement, which are often available. Where data are not available, EPA may make casespecific assumptions based on cultural practices and physical characteristics of the crop plants. 

Plant material containing dsRNA would act as a source of exposure for organisms that directly 

ingest plant tissue; however, once plant tissue breaks down and cells lyse, the dsRNA may also 

be released into the environment. Degradation of the plant tissue will affect the potential for 

exposure, since RNA is degraded within the plant during senescence (Pietramellara et al., 2009). 

However, not all dsRNA is necessarily expected to be degraded by the time plant material 

reaches the ground. Once released from plant cells, dsRNA is expected to ultimately end up in 

the soil in terrestrial environments, where it may persist or be transported to other terrestrial or to 

15
aquatic areas. Plant material that is moved into aquatic areas would provide a source of dsRNA 

in aquatic environments, where it may move into sediment or be present in the water column. 

Studies show that DNA can persist in soil by binding to humic substances or minerals, or it may 

be degraded by microbes and nucleases (Levy-Booth et al., 2007; Pietramellara et al., 2009). 

DNA can persist for a few days in soil or for several years (Levy-Booth et al. 2007). Little is 

known about the fate of dsRNA in the environment, and the chemical and structural 

characteristics of RNA and DNA may be sufficiently different to prevent use of data developed 

on the environmental fate of DNA as a surrogate for RNA. In the absence of any environmental 

fate data, EPA can estimate conservative environmental exposure concentrations based on 

expression data and assumptions about persistence; however, EPA typically receives soil 

degradation data on Bt-derived PIPs, which provide information about the potential for their 

persistence in the soil. Thus far, these studies, in addition to longer term (i.e. multi-year) soil 

degradation studies, have been considered to be sufficient in developing an understanding of the 

persistence of Bt-derived PIPs in the environment. While focus only on soil degradation may

also be adequate for dsRNA PIPs, additional information on the potential for dsRNA PIPs to 

move vertically and laterally with water in soil, bind to soil and sediment particles, and the 

effects of binding to soil particles on persistence and bioavailability (e.g., by protection from 

nucleases, see Pietramellara et al. 2009) could aid in determining the spatial extent of exposure 

and the on- and off-field levels of exposure to nontarget organisms. 

b. Nontarget Exposure Opportunities

In risk assessments for PIPs, EPA typically considers exposure to birds, wild mammals, 

freshwater and marine/estuarine fish, aquatic invertebrates, nontarget insects, including honey 

bees, nontarget plants, and soil invertebrates. These taxa are considered sufficient representatives 

of the potential for risk to all nontarget organisms for purposes of screening level risk 

assessments, though refined risk assessments may consider exposure at lower taxonomic levels.

For most PIPs, since they are expressed in the crop plant, nontarget organism exposure is 

expected to occur primarily through ingestion of the living or dead plant material, which can 

occur on or off the field in which the crop is growing. Nontarget organisms, particularly insects, 

may consume pollen produced by the crop plant, either while it is on the plant or once it has 

fallen within the crop field or outside it. Pollen may be consumed directly as food (or, in the 

case of bees, may be carried back to the hive or nest to feed to larvae), or may be consumed 

incidentally on food plants upon which the pollen falls. While pollen consumption may be more 

likely to be a means of exposure for insects, it is also possible for other animals (e.g., grazing 

vertebrates) to consume plants upon which pollen has fallen. Plant material that becomes dust, as 

described above, could also incidentally expose nontarget organisms to dsRNAs. 

16
Secondary exposure may also occur, where organisms that have ingested dsRNA in plant tissue 

(e.g., nonsusceptible pests) are consumed by nontarget predators or parasites. The factors 

influencing the possibility of exposure by this pathway (e.g., longevity of dsRNA once 

consumed, concentration resulting within the herbivorous insect) are not known. However, 

Whyard et al. (2009) observed susceptible insects feeding on dsRNA for several days before 

deleterious effects were observed, which would present an opportunity for this type of exposure. 

Garbian et al. (2012) also showed that dsRNA could be transferred from bees to Varroa mites, 

indicating that this type of exposure is possible. 

Ingestion of pesticidal dsRNA that is free in the environment (following plant tissue breakdown) 

may also occur if it is present in soil, sediment or water. Incidental ingestion of dsRNA present

in soil and sediments by nontarget organisms is a possibility. This is expected particularly of 

invertebrates that live in soil or sediment; however, incidental soil ingestion by vertebrates 

foraging along these substrates could also occur (e.g., birds probing soil for invertebrates). Any 

free dsRNA in the water column may be inadvertently consumed by both vertebrates and 

invertebrates, both by organisms living in such environments and those organisms that use them 

as a source for water. 

Exposure to free dsRNA via the integument (in animals) is not a route that has been considered 

in nontarget risk assessments for Bt-derived PIP proteins; however, evidence exists that this may 

be a possibility for dsRNA. Whyard and colleagues cite examples in which RNAi was induced in 

nematodes and flatworms by soaking these animals in a solution of dsRNA. They also cite 

research in which gene silencing was observed in dechorionated Drosophila embryos (see 

Tabara et al., 1998; Orii et al., 2003; and Eaton et al., 2002 as cited in Whyard et al., 2009).

Campbell et al. (2010) also observed significant gene knockdown in Varroa destructor with 

soaking in a solution of NaCl and dsRNA. More information is needed, particularly since the 

method of soaking in these experiments could have actually led to ingestion of the dsRNA, but 

these data may indicate the possibility of exposure through this route. Organisms that might be 

exposed in this way could include animals in aquatic environments as well as those inhabiting 

soil and sediment, though EPA recognizes that the exposure concentration in these environments 

may be low. Uptake by plants in both terrestrial and aquatic environments may also be possible, 

but little information also exists on the potential for plant exposure to dsRNAs in the 

environment. 

The above is meant to provide a comprehensive listing, in light of the limited information 

available on dsRNA in the environment, of potential routes through which nontarget organisms

might be exposed to dsRNAs expressed in crop plants. EPA recognizes that not all of the routes

discussed above may be significant to the risk assessment, since some may have low likelihood 

of occurring and other factors (e.g., environmental concentration, barriers to uptake) may also 

limit exposure. Nonetheless, at this point in time, recognition of the potential of these scenarios 

17
to affect nontarget organisms might be advisable in assessing the risk of dsRNA PIPs, even if 

only to show that these scenarios are not a consideration for the particular PIP. 

c. Other Factors Affecting Exposure Levels

The level and duration of exposure can be influenced by additional factors, even once dsRNA 

exposure has occurred. Exposure by ingestion to dsRNA may be reduced by barriers that exist in 

the gut. Potential barriers for uptake in mammals were discussed previously for human health 

exposure and effects. Some evidence for similar barriers also exists for arthropods. For example, 

Allen and Walker (2012), observed successful gene knockdown in the tarnished plantbug (Lygus 

lineolaris) when dsRNA was injected into the hemocoel; however, when dsRNA was fed to the 

insect, RNAi was not successful. The lack of effects was attributed to dsRNA digestion by 

nucleases in the insect gut. This study also showed that the saliva of L. lineolaris degraded 

double-stranded RNA. On the other hand, exposure by both injection and feeding successfully 

resulted in reduced expression of targeted genes in the triatomine bug, Rhodnius prolixus (Araujo 

et al., 2006). Other conditions of the gut environment, such as pH, may also influence breakdown 

of dsRNA, and mechanisms of uptake may also vary (Terenius et al., 2011; Katoch et al., 2013). 

However, much of what we understand about RNAi in living organisms is limited to mammals 

and arthropods, and little work has focused on the barriers to uptake that exist in other 

organisms. Better understanding of the mechanisms influencing uptake, particularly if they can 

be extrapolated to other organisms, would reduce uncertainty in exposure assumptions and help 

to focus risk assessments on the most appropriate organisms. 

For purposes of assessing risk, exposure levels are typically assumed to be equivalent to the 

highest amount expressed among various plant tissues, except where the focus may be exposure 

to a particular type of plant tissue (e.g., exposure via pollen collected by bees). The effects of 

dsRNA are understood to be dose-dependent, and for some organisms, exposure must be 

continuous to result in effective gene silencing (Katoch et al. 2013). Evidence also exists of the 

potential for upper and lower thresholds to the effects of gene silencing. As discussed previously, 

mammalian cell culture studies indicate that a minimum number of copies of siRNA may be 

needed at a targeted site to induce RNAi, whereas Fire et al. (1998) suggest that only a few RNA 

molecules are sufficient to initiate the RNAi pathway. Huvenne and Smagghe (2009) also 

suggest that an optimum concentration exists for gene silencing for each target gene and 

organism, and that exceeding this optimum does not necessarily result in additional gene 

silencing. Based on this evidence, there may be environmental exposure levels below which no 

effects, or above which no greater degree of effects, would necessarily occur. It is also possible 

that lower limits of exposure may be essentially non-existent for some species. More clarity on 

this issue and its potential influence on exposure estimates are needed.

18
Some organisms have the ability to amplify the RNAi signal by utilizing environmental siRNA 

as a template to produce additional siRNA, leading to exposure within the organism that is more 

prolonged and greater than that taken up. siRNA amplification has been observed in C. elegans, 

which accomplish this through RNA-dependent RNA polymerase (RdRP)(Sijen et al., 2001). 

This phenomenon has also been observed in plants and fungi (Dillin, 2003). While amplification 

by RdRP has been characterized in these organisms, whether a similar mechanism exists in other 

organisms has not been determined. If the ability to amplify the RNAi signal is possible in other 

organisms, this may affect the usefulness of exposure estimates involving those organisms, since 

environmental exposure determined to be “low” may not translate to the same exposure within 

the organism. On the other hand, if uptake at the cellular level is not possible in some organisms, 

then the environmental exposure would be irrelevant. Further understanding of these 

mechanisms would aid in deriving exposure estimates and would reduce uncertainty.

2. Ecological Effects to Nontarget Organisms

a. EPA’s Current Testing Framework

Data requirements for PIPs are determined on a case-by-case basis, and are based primarily on 

the tiered microbial pesticide data requirements for environmental fate and nontarget effects in 

40 CFR 158.2150. The list below describes what are considered Tier I nontarget effects and 

environmental fate studies that are typically received for new PIP active ingredients. These 

studies generally have been considered sufficient to inform nontarget risk assessments for Btderived PIPs to date. The preferred test material for these studies is the Bt protein, extracted from 

a microbial source (e.g., Escherichia coli) and purified; however, in some cases, plant material 

(e.g., grain, pollen, lyophilized leaf tissue) has also been used. In these studies, test subjects are 

typically fed a maximum hazard or limit dose, and mortality is the endpoint measured. Some 

studies with insects (historically the focus of Bt-derived protein PIP nontarget risk assessments) 

have also included observations of effects on growth and reproduction of test subjects. Studies 

that are considered higher tier studies, such as field studies on the effects of beneficial insects or 

multi-year soil persistence studies, have also been required, though they are currently not relied 

upon as heavily for new Bt-derived PIP active ingredients. 

Nontarget effects

Wild mammal toxicity Honey bee toxicity (adult and larvae)

Avian oral toxicity Nontarget plant toxicity

Avian dietary toxicity Arthropod toxicity

Freshwater fish toxicity Non-arthropod invertebrate toxicity

Freshwater invertebrate toxicity Monarch butterfly (in specific cases) 

Estuarine/marine animal toxicity Synergistic effects (for combination PIPs)

Environmental Fate

19
Soil degradation

b. Unique Challenges Presented by dsRNA PIPs 

Risks of deleterious effects of Bt-derived PIPs on nontarget organisms assessed by EPA thus far 

have been determined to be minimal, and this is due in large part to the well-characterized 

specificity of Bt proteins. The current testing framework for PIPs, relying primarily upon first 

tier tests with focus on acute endpoints, is considered to be adequate for informing nontarget risk 

assessment for these types of PIPs. While dsRNA produced in crop plants as a means of pest 

control has been claimed to have a high degree of specificity (Petrick et al. 2013; Whyard et al. 

2009), evidence suggests that it may have other unexpected effects. Therefore, dsRNA PIPs may 

present new challenges for ecological risk assessment.

In general, specificity does not appear to be guaranteed for dsRNAs when fed to insects. For 

example, Whyard et al. (2009) observed slight cross-species silencing among four insect species 

fed dsRNAs that were meant to target genes in other species. Terenius et al. (2011) also were not 

able to confirm specificity of RNAi following examination of data from numerous studies with 

RNAi in insects, and also proposed that variation in sensitivity of the target gene to RNAi and in 

the sensitivity of the tissue targeted may influence effects of dsRNA. Baum et al. (2007) 

observed target gene silencing and reduced survival in Leptinotarsa decemlineata, but silencing 

also occurred in an ortholog to the target gene in Diabrotica virgifera virgifera, although 

silencing in the latter species required exposure to greater dsRNA concentration. Therefore, even 

when intended to target a specific gene, effects among related organisms have been observed. 

“Off-target” effects may also result from binding of siRNAs to genes other than the target. 

These effects appear to be related to partial sequence homology between the “seed region” 

(positions 2-7 or 2-8) of the siRNA and the 3’ untranslated region of messenger RNA transcribed 

from a nontarget gene (Birmingham et al. 2006, Jackson et al. 2003), though Birmingham et al.

(2006) note that such binding is likely one factor in a complex process that is not yet fully 

understood. Off-target effects are siRNA specific, instead of target-gene specific, and may 

influence other signaling and transcription pathways (Jackson et al. 2003). They have also been 

shown to reduce cell viability in controlled studies (Fedorov et al. 2006). Since RNAi reduces or 

eliminates translation of a mRNA gene product into protein, but does not alter gene expression 

directly (Huvenne and Smagghe 2010), off-target effects may not be severe or prolonged if they 

were to occur. This will, of course, depend on the sensitivity of the nontarget organism, the 

gene(s) targeted, and the potential for amplification of siRNAs. Nonetheless, little information is 

available with which to estimate the likelihood of these effects resulting from unpredicted 

interactions of dsRNA with genes not intended for silencing and they may be significant factors 

affecting nontarget risk (Auer and Frederick 2009, Lundgren and Duan 2013). 

20
Systemic RNAi, discussed in more depth in section II.C., could be an additional contributing 

factor in the observation of unexpected effects. For example, Terenius et al. (2011) describe 

work with Hyalophora cecropia in which RNAi was observed in the offspring of insects injected 

as pupae, indicating uptake of the dsRNA by oocytes in the pupae. Such effects also indicate a 

potential influence of life stage on RNAi, which is discussed further below. On the other hand, 

Whyard et al. (2009) failed to observe systemic RNAi in D. melanogaster; therefore, systemic 

RNAi may be possible in some species but not others. The mechanisms driving the systemic 

RNAi among species are not known, and studies to date do not cover the possibility of this 

effect, or lack thereof, in all taxa that EPA considers in its risk assessments. 

RNAi has necessary functions within cells that are important to growth, development and tissue 

homeostasis. Oversaturation of RNAi machinery as a result of introduction of environmental 

dsRNA could disrupt regulation of gene expression and normal cell function (Dillin 2003, 

Katoch et al. 2013, Lundgren and Duan 2013). Saturation could also lead to reduced defenses 

against viral infection (Dillin 2003). Exposure to RNAi may also stimulate immune response. 

This has been observed in mammals (see Lundgren and Duan 2013), and could be a factor for 

observed gene silencing in some insect studies (Terenius et al. 2011); however, how immune 

stimulation by RNAi may affect nontarget organisms is not known. 

c. Possible Limitations of the Current PIP Testing Approach

Considering the unique mode of action of dsRNAs that may serve as PIPs and the unintended 

effects of RNAi gene silencing described above, it would seem that risk assessors should remain 

cognizant of the possibility of unexpected effects that may not be readily predicted based on the 

known specificity for the target gene. The current approach for testing/assessing PIPs for 

potential risk will serve to provide much information for risk assessment; however, owing to the 

nature of dsRNA and RNAi, some questions may remain as to how well the current suite of 

required studies can detect potential effects of dsRNA PIPs on nontarget organisms. 

Better information with which to estimate exposure could be valuable to ecological risk 

assessments for dsRNA PIPs. Some data are available with which to determine the potential for 

dsRNA uptake in mammals, which may be extrapolated to other vertebrates with similar 

physiological characteristics (e.g., acid stomachs); however, less information is available for 

vertebrates with other characteristics. Additionally, some research has focused on barriers to 

uptake in arthropods, but clear trends are not available with which to make predictions. More 

information on barriers to dsRNA uptake or dsRNA degradation after uptake, particularly data 

that would facilitate predictions across nontarget taxa, could refine exposure estimates or allow 

assumptions of minimal exposure in certain organisms.

21
More information on specificity and the realistic possibility and impacts of the unintended 

effects described above on nontarget organisms may also help to focus testing needs. At this 

point, unintended effects primarily have been described from laboratory screening tests (e.g., 

microarray analyses) or testing in which the exposure was artificial (e.g., microinjection in 

insects). Without further information, it is not unreasonable to assume that some kind of related 

effect could occur in nontarget organisms; however, at this point the actual biological impacts 

are not known. 

EPA would rely on environmental fate and nontarget toxicity testing to inform the ecological risk 

assessment; however, the unique nature of dsRNA and RNAi raise several issues of concern with 

respect to the typical data set submitted for nontarget effects: 

1) The potential influence of latent effects on results of nontarget testing. As discussed above, 

arthropods in laboratory studies have been observed feeding for several days on dsRNA before 

the effects of gene silencing were apparent. Several of the toxicity studies submitted for PIPs are 

short term dosing studies, and latency of effects may impact observations of both sublethal 

effects and mortality. Some studies, such as nontarget insect studies, are carried out for sufficient 

time to observe effects on reproduction, and latent effects would more likely be observed. 

Therefore, the importance of this issue could vary between different types of studies.

2) The appropriate life stage for testing. Toxicity testing is often targeted toward young test 

organisms, as it is assumed that younger organisms are more sensitive to toxic substances than 

adults. Much of the work that has been done with arthropods suggests that younger life stages are 

more sensitive to RNAi (Huvenne and Smagghe, 2009). However, sensitivity may not follow a 

specific trend over time, but may be the result of the importance of a target gene, or a gene 

affected by off-target silencing, in the organism at a specific life stage. It is likely that dsRNA 

designed for delivery to pest insects in plants would target young life stages, so testing on those 

life stages would be appropriate. However, given the range of possible unexpected effects, it is 

conceivable that an effect could occur in the field that would not be observed in the lab. 

3) The possibility of chronic effects. Current testing focuses on mortality as the endpoint. Offtarget effects, or binding to target genes in a less sensitive organism, could cause effects that do 

not lead to lethality. This has been observed in studies with insects. For example, Zha et al. 

(2011) observed suppression of target genes encoding a hexose transporter, a carboxypeptidase, 

and a serine protease in the brown planthopper (Nilaparvata lugens Stål), but did not observe 

significant lethality. Suppression of genes without overt signs of toxicity may be considered 

insignificant following a single exposure; however, long-term exposure and continuous or 

repeated gene knockdown could result in chronic effects. Since pesticidal substances are 

produced continuously in a PIP crop plant, chronic exposure to some organisms is possible. 

22
Approaches other than nontarget testing may provide additional information and reduce data 

requirements. For example, while there are currently limitations to the use of bioinformatics

analysis, particularly for nontarget organisms for which there are limited genomic data, this type 

of approach may be useful in identifying groups of nontarget organisms that are more or less 

susceptible to the effects of dsRNA PIPs. Further development of predictive tools also has 

potential to reduce uncertainty as well as the scope of testing required to support a risk 

assessment. 

3. Conclusions – dsRNAs Expressed As PIPs

RNAi gene silencing is a relatively recent discovery, and there are general uncertainties with the 

current state of knowledge about dsRNAs and their potential environmental fate, as well as the 

importance of effects observed in laboratory studies to actual nontarget hazard. The current 

approach as used for Bt-derived PIPs may be sufficient; however, information in the literature 

suggests the need to consider additional effects beyond those typically investigated for Btderived PIPs. At present, EPA could move forward with a risk assessment based on the current 

approach; however, the resulting risk predictions could have uncertainty or require conservative 

assumptions. Therefore, EPA seeks guidance on the natural processes in the environment and 

within nontarget organisms that serve to reduce or eliminate exposure, the importance of the 

potential for unexpected effects of dsRNA PIPs and RNAi in nontarget organisms, and additional 

testing that EPA may require to reduce uncertainties in risk estimates. 

B. Non-PIP dsRNA End Use Products

The current approach used by the Agency to assess the ecological risks of traditional chemical 

pesticides (conventional chemicals and biochemicals) may also be used as a basis for ecological 

risk assessment of non-PIP dsRNA-containing end-use products (dsRNA-EPs). As stated above 

in Section IV of this paper (Environmental Considerations for PIP-expressed dsRNA), 

environmental fate and effects of dsRNA are poorly understood and present unique challenges 

for ecological risk assessment that have not yet been encountered in assessments for traditional 

chemical pesticides 

The Agency has many years of ecological risk assessment experience with PIPs and the previous 

section of this document has explored the various pathways whereby dsRNA expressed in PIPs 

may move into the environment and result in nontarget exposure. Unlike PIPs, however, the 

Agency has not, to date, assessed the hazards or risks of dsRNA applied directly to the 

environment as components of end-use products intended for pest control under Section 3 of 

FIFRA. Numerous ecological risk assessments have been conducted on traditional chemical 

pesticides (EPA, 2012a). However, the screening level assessments currently used for traditional 

23
chemical pesticides may not be applicable due to the unique modes of action of dsRNA active 

ingredients. As stated above in Section IV.A., EPA’s goals are to ensure that unreasonable 

effects do not occur to nontarget populations. Details of the approaches used by Office of 

Pesticide Programs to conduct ecological risk assessments on traditional chemical pesticides may 

be found at EPA’s Ecological Risk Assessments webpage (EPA, 2012b).

This section of the paper will focus on descriptions of the various exposure pathways that may 

result following the application of dsRNA-EPs. The potential human health and 

ecotoxicological effects of dsRNA have been discussed in detail in previous sections of this 

paper, and will be discussed here only in the context of uncertainties and likely data needs. In 

order to delineate the possible pathways of exposure, there first will be a discussion of the likely 

categories of dsRNA active ingredients intended for use in pest control products, as well as the 

likely end-use products and application methods. This section of the paper will conclude with a 

brief discussion of issues and uncertainties related to estimation of environmental exposure and 

determination of effects, the two major components of an ecological risk assessment, for non-PIP 

dsRNA-EPs. 

At the time of this writing, no entity has approached the Agency with an application to register a 

non-PIP dsRNA-EP. While based on publically-available information, the following discussion 

regarding the different categories of dsRNA active ingredients and application methods, is 

speculative. 

1. Likely Categories of Active Ingredients

At his time, the Agency anticipates four likely categories of dsRNA active ingredients that could 

be present in non-PIP dsRNA-EPs: (1) Direct control agents; (2) Resistance factor repressors; 

(3) Developmental disruptors; and (4) Growth enhancers.

a. Direct Control Agents

A dsRNA direct control agent is defined here as a dsRNA active ingredient that has direct toxic 

effects upon the metabolism of the pest resulting in mortality. The difference between dsRNA 

activity and traditional chemical pesticides is that dsRNA operates at the level of the gene, 

whereas traditional pesticides have toxic physiological effects at higher levels of organismal 

metabolism. The family of dsRNA direct control agents likely would include, but are not limited 

to herbicides, insecticides, and fungicides. This dsRNA active ingredient category does not 

depend upon chemical pesticide control, and could be rotated into integrated pest management 

(IPM) systems to reduce chemical pesticide use and lessen the possibility of resistance 

development by the target pests. 

24
b. Resistance Factor Suppressors

A dsRNA resistance factor suppressor is defined here as a dsRNA active ingredient that 

suppresses genetic resistance to a traditional chemical control. Many pest species have become 

resistant to chemical control since the advent of widespread chemical pesticide use in the 1940s 

and there is a long history of the development of resistance to chemical controls by insects 

(IRAC, 2013). More recently, herbicide resistant weeds have becoming a major worldwide pest 

control problem (Mortensen et. al., 2012). As the list of pesticide-resistant insect and weed 

species grows longer, there is a proportional interest in developing dsRNA products that 

specifically target the genes for resistance to specific herbicides and insecticides (Lundgren and 

Duan, 2013; Preuss and Pikaard, 2003). This dsRNA active ingredient category does not reduce 

dependence upon chemical pesticide control, but does permit the continued use of existing 

chemistry by rendering formerly resistant pests susceptible. 

c. Developmental Disruptors (Growth Regulators)

A dsRNA developmental disruptor is defined here as a dsRNA active ingredient that interferes 

with the normal development or growth of the target pest such that the target pest or its progeny 

either die (indirect mortality), are less competitive, or are sterile. Developmental disruptors that 

are currently registered by the Agency are insect growth regulators (IGRs), and they fall into two 

main categories: (1) juvenile hormone mimics (juvenoids) that disrupt hormonal control of 

larval development and inhibit metamorphosis (e.g. methoprene); and (2) chitin synthesis 

inhibitors (e.g. triflumuron) that prevent chitin formation and replacement of the old cuticle 

following ecdysis (molting). It is conceivable that dsRNA active ingredients could be developed 

that specifically target genes responsible for the proper growth and development of any taxa of 

pest species, not just that of insects. 

d. Growth Enhancers

A dsRNA growth enhancer is defined here as a dsRNA active ingredient that stimulates, inhibits, 

or otherwise mimics the activity of a naturally-occurring plant hormone. Plant hormones and 

their analogues are considered pesticides under FIFRA (1996) and are regulated as such. Plant 

hormones are typically applied to food and ornamental crops for aesthetic purposes (color, odor, 

taste, shape, uniformity), as harvest controls (delayed or advanced maturity, inhibition or 

promotion of fruit ripening, inhibition or promotion of abscission), and for resistance to 

environmental stress. Another category of plant regulators not associated with the traditional 

plant hormones, the induced resistance promoters, are those substances that stimulate the internal 

defense mechanisms of plants such that they will have an enhanced capacity to resist infection by 

plant pathogens. The two most well-known mechanisms of induced resistance are known as 

Systemic Acquired Resistance (SAR) and Induced Systemic Resistance (ISR) and they are 

25
distinguished from one another primarily by how the respective resistance mechanisms are 

elicited, the subsequent pathways of resistance induction, and the specificity of the induced 

response (see review by Jones, 2012). It is conceivable that dsRNA active ingredients could be 

developed that specifically target genes responsible for pathogen resistance. This type of dsRNA 

product could be used in two ways: (1) stimulate pathogen resistance in desirable food and 

ornamental plants; and (2) suppress pathogen resistance in weed species. In addition, it is 

conceivable that a family of dsRNA products could be developed for the purpose of suppressing 

disease resistance in other pest taxa (e.g. birds, fish, insects, mammals etc.)

2. Potential Application Methods

It is anticipated that dsRNA-containing end-use products could be applied using the same 

methods as traditional chemical pesticides:

• Foliar sprays

• Post-emergence directed soil spray/drench

• Seed treatments (as a protectant)

• Granular/powder (soil applied topical/incorporated & baits/traps) 

Depending upon the target pest, application method, and product type, both outdoor and indoor 

use sites also are anticipated. Outdoor use sites may include, but are not limited to field crops, 

nurseries, forests and other natural areas; outdoor use sites also may include both terrestrial and 

aquatic habitats. Indoor use sites may include, but are not limited to food storage facilities, 

greenhouses, and residential/structural treatments. 

3. Nontarget Exposure 

As stated above in Section IV.A., the degree of nontarget exposure depends on the distribution 

and fate of dsRNA within the environment, as well as the potential routes of direct and indirect 

exposure, exposure duration, and the potential for uptake by nontarget species. In regard to nonPIP dsRNA EPs, the distribution and fate of dsRNA within the environment, potential routes of 

direct and indirect exposure, and duration of exposure to nontarget organisms will be dependent 

upon the following: 

• Application rate of active ingredient

• Application timing

• Application method

• Number of applications

• Off-site movement of applied dsRNA

• Stability and persistence of exogenously applied dsRNA following application 

26
Once the product has been applied, other factors will affect the distribution of dsRNA in the 

environment and the potential for direct and indirect exposure to nontarget organisms. The most 

important of these factors are: (1) In situ dissipation of dsRNA via abiotic (e.g. UV-light) and 

biotic (microbial degradation) processes; (2) Offsite (physical) movement of applied dsRNA 

from the treatment area; and (3) persistence of dsRNA in the environment. 

Stabilized vs. Unstabilized dsRNA. The following discussion regarding the factors influencing 

the distribution and persistence of dsRNA in the environment presumes that exogenously applied 

dsRNA will be stabilized in some manner to inhibit abiotic and biotic degradation in the 

environment such that there will be sufficient residence time in/on the treated use site to permit 

the maximum desired pesticidal activity. Similarly, for products requiring ingestion by the pest 

(e.g. treated foliar surfaces, granules, etc.) the active ingredient likely will be stabilized for a 

sufficient period of time for the target pest to locate the treated article and consume a sufficient 

amount of dsRNA to have the desired pesticidal effect. It is further presumed that the 

degradation rate of “stabilized” dsRNA will likely be slower than “unstabilized” dsRNA. 

a. In situ Dissipation of dsRNA

It is presumed that once applied at the use site, dsRNA will be subject to the same abiotic and 

biotic degradation processes as traditional chemical pesticides. However, as described above in 

Section IV.A., few data are available regarding the degradation of dsRNA in the environment 

(see review by Lundgren and Duan, 2013). Data regarding the fate of DNA in soil and sediment 

(Levy-Booth et. al, 2007; Pietramellara et al. 2009) may not be applicable to understanding the 

fate of dsRNA due to the different chemical characteristics of the two molecules. Available data 

indicate that RNA may degrade more rapidly than DNA, although DNA in soil may degrade in a 

little as 7 days or persist for several years, with persistence enhanced by binding to soil humic 

substances, or degraded rapidly by microbial activity (Levy-Booth et. al, 2007 as cited in 

Lundgren and Duan, 2013). The relative degradation rates of DNA vs. RNA in the environment 

remains nebulous. 

There also is uncertainty regarding whether exogenously applied dsRNA will amplify within 

living plant tissue once it has been absorbed and if so, the degree of amplification that will occur 

prior to loss of the tissue by the plant. While dsRNA present on treated plant surface may begin 

to degrade, it is possible that amplification within plant tissues, if it occurs to any substantial 

degree, may result in tissue residue levels that are greater than at the time of application, further 

complicating EEC determinations. 

Another complicating factor is the degradation rate of “stabilized” dsRNA vs. “unstabilized” 

dsRNA. The Agency presumes that stabilized dsRNA will degrade at a slower rate in soil and 

water, and in/on biological matrices than unstabilized dsRNA-EP. 

27
b. Off-site Movement of Applied dsRNA

Offsite movement of dsRNA from the treatment site may occur via the following pathways:

• Surface runoff from foliage and soil following a precipitation event

• Infiltration into the soil and movement into groundwater

• Spray drift

• Physical movement of treated plant tissue and pollen 

• Pollinators and other animal vectors. 

Regardless of whether the dsRNA is unstabilized or stabilized it will be distributed throughout 

the environment via the pathways described above. Volatilization into the atmosphere is not 

expected. The first three pathways listed above are self-explanatory and indicate how dsRNA 

may migrate from the application site to offsite terrestrial and aquatic environments. Plant tissue 

and pollen movement were discussed above in Section IV and would raise the same exposure 

concerns if exogenously applied dsRNA is amplified within treated plant tissues and transported 

into, or expressed in pollen, or if dsRNA was applied during anthesis. 

Pollinators such as honey bees, if present on plants at the time of application, may carry dsRNA 

residues back to their hive and distribute the residues throughout the hive via bee-to-bee contact. 

Similarly, dsRNA residues could be carried back to the hive in pollen if exogenously applied 

dsRNA is amplified in treated plant and expressed in the pollen. Although this may not be a 

major offsite pathway from the treated area, dsRNA residues could bioaccumulate within the 

hive and have unanticipated effects resulting from off target effects. Bees can also transfer 

dsRNA to other organisms. Garbian et. al. (2012) demonstrated that dsRNA ingested by bees 

could be transferred to Varroa mites ( a pest of bees) and then from the mite to a Varroa-infested 

bee. Other horizontal transfers may be possible.

Migration of dsRNA from the application site may occur via other animal vectors. Random 

movements of insects, as well as terrestrial and avian wildlife through treated areas at, or 

following the time of dsRNA application, could carry unknown amounts of dsRNA to offsite 

areas although this is not likely to be a major offsite pathway and would be difficult to model. 

c. Persistence of dsRNA in the Environment

Accurate determination of exposure, hazards, and risk of exogenously applied dsRNA requires a 

complete understanding of dsRNA persistence, or lack thereof, in the environment. At this time, 

there are no quantitative measures of dsRNA degradation in soil, water or biological matrices, 

under aerobic and anaerobic conditions, or under conditions of variable pH and UV light. 

28
Modeling of Environmental Fate. There are a number of models developed by the EPA to 

estimate the environmental concentrations of chemical pesticides, following applications at 

maximum label use rates, for use in estimating pesticide exposure to non-target organisms (EPA, 

2013). Unfortunately, these models are of limited utility for determining the estimated 

environmental concentrations (EECs) of exogenously applied dsRNA, due to the paucity of 

information regarding degradation rates of dsRNA in soil and water matrices and under aerobic 

and anaerobic conditions. Furthermore, it is unknown whether exogenously applied dsRNA will 

amplify within living plant tissue once it has been absorbed and if so, the degree of amplification 

that will occur prior to loss of the tissue by the plant. In situ amplification of dsRNA in plant 

tissue would complicate environmental exposure estimations calculated by the existing models. 

4. Hazards to Nontarget Organisms and Environmental Fate

At this time, the Agency does not possess empirical data regarding the acute or dietary hazards 

of dsRNA to non-target birds, fish, aquatic invertebrates, insects, and plants. However, the 

Agency believes that the existing hazard and risk assessment paradigm of tiered toxicity testing 

for biopesticides may be applied to assess the environmental fate and effects of dsRNA. Specific 

data requirements have been developed by the Agency to assess the hazard to nontarget 

organisms from exposure to new active ingredients. The list below describes the Tier I nontarget 

organism and environmental studies that must be addressed to support the registration of a new 

biochemical pesticide active ingredients and are similar to those used to determine the acute 

hazards of microbial pesticides. These studies have been generally been considered sufficient to 

inform nontarget risk assessments for all new biochemical pesticides to date. The preferred test 

material is the technical grade active ingredient (TGAI), the product that is the purest and highest 

concentration of the active ingredient that is typically used to formulate one or more end-use 

products. The studies focus primarily on maximum hazard dose testing and use mortality as a 

toxicological endpoint. The Tier I biochemical pesticide nontarget organism and environmental 

fate data requirements may be found in the table at 40 CFR 158.2060(d) and are summarized 

below:

• Avian acute oral toxicity

• Avian dietary toxicity

• Fish acute toxicity, freshwater

• Aquatic invertebrate acute toxicity, freshwater

• Terrestrial plant toxicity, seedling emergence

• Terrestrial plant toxicity, vegetative vigor

• Non-target insect testing

29
As discussed in Section IV.B.3. above, the persistence of dsRNA in the environment is 

relatively unknown. Although rough estimations of degradation rates may be calculated using 

the limited data available for the environmental fate of similar nucleic acids, specific information 

is lacking on dsRNA, regardless of whether it is stabilized when formulated into end-use 

products. The Agency believes that selected Tier II biochemical pesticide environmental fate 

studies can be used to determine the degradation rate and environmental fate of dsRNA 

following application of dsRNA-EPs to plant and soil surfaces. Data obtained from these studies 

will more completely inform exposure assessments for exogenously applied dsRNA products, as 

well as for dsRNA-PIPs. The Tier II biochemical pesticide nontarget organism and 

environmental fate data requirements may be found in the table at 40 CFR 158.2060(d) and are 

summarized below:

• Sediment and soil adsorption/desorption for parent and degradates

• Soil column leaching

• Hydrolysis

• Aerobic soil metabolism

• Photodegradation in water

• Photodegradation on soil

• Anaerobic soil metabolism

• Aerobic aquatic metabolism

• Anaerobic aquatic metabolism

All of the studies listed above must be supported with robust and independently validated 

analytical methods that are accurate and precise for the detection of dsRNA residues in various 

environmental matrices. 

5. Possible Limitations of the Current Testing Paradigm for Biochemical Pesticides

Biochemical pesticides are naturally-occurring substances that have non-toxic, wellcharacterized modes of action against their respective target pests. Acute and subchronic hazard 

data, if not readily available from other sources, are typically easy to generate using the Tier I 

studies listed above in Section IV.B.3. Acute and subchronic hazards of dsRNA to nontargets 

may likewise be determined using the same battery of testing. The new categories of dsRNA 

products, however, will present additional hazard and risk assessment challenges due to their 

unique modes of action and other toxicological endpoints that cannot be measured using the 

traditional testing paradigm. In a recent review, Lundgren and Duan (2013) observed that the 

current tiered hazard assessment approach used by the Agency, is inappropriate to address the 

following unique hazards potentially posed by dsRNA products:

30
• Off-target gene silencing

• Silencing the target gene in unintended organisms

• Immune stimulation

• Saturation of the RNAi machinery in cells

All of the above potential dsRNA hazards have been discussed in detail in previous sections of 

this document and will not be revisited in this section of the document; other unanticipated 

dsRNA hazards also may occur. The Agency assumes that the hazards and risks to nontargets 

identified in Section IV.A., will be identical for non-PIP dsRNA-EPs, with the exception that the 

exposure suite will be much greater for exogenously applied products. We note that accurate, 

standardized methods for measuring and assessing the aforementioned hazards will be necessary 

to conduct robust nontarget species risk assessments on dsRNA products. 

6. Conclusions

Non-PIP dsRNA end-use products (dsRNA-EPs) present new challenges in regard to ecological 

risk assessment for nontarget taxa. These challenges arise as a result of the unique modes of 

action of dsRNA active ingredients,and the paucity of reliable data regarding the persistence and 

environmental fate of exogenously applied dsRNA. Persistence and environmental fate will 

strongly influence the degree of exposure to nontarget organisms, nontarget effects, and final 

determination of risk. In regard to hazard effect, the current testing paradigm for nontarget 

species characterizations, which emphasizes limit dose testing and use of mortality as an 

endpoint, likely will not be adequate to assess adverse effects resulting from off-target gene 

silencing, silencing of the target gene in unintended organisms, immune stimulation, and 

saturation of the RNAi machinery in cells. 

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