2008-09 Gathering of Voices

Phytotoxic effects of the environmental endocrine disruptor bisphenol A on Brassica Rapa (Wisconsin fast-start® mustard plants)

Evan Knappenberger

Abstract

Bisphenol A (BPA), an environmental endocrine disruptor, (xenoestrogen), is a common environmental pollutant as well as a byproduct of plastic and resin manufacturing. As a contaminant, it can be found in household products, sewage sludge, watersheds and many types of plastic consumer products. Studies have documented its clastogenic, teratogenic, carcinogenic and pathological effects on both plants and animals, and have demonstrated its sorption ability into soils. It has been suggested that BPA accumulates in agricultural crops and edible plants, thus entering the food chain and posing a significant risk to human consumption. In this experiment, Brassica Rapa (Wisconsin Fast-Start © Mustard) was irrigated with treatments of 10 mg/L and 50 mg/L BPA for 26 days including germination and flowering periods. Evidence of phytotoxicity was apparent at the higher dose of 50 mg/L, including disrupted growth patterns and sexual maturation over the course of the 30-day lifespan. Effects were measured at 4, 8, 10, 15, 19, 23, and 26 days. Generally, deformities and survivability of Brassica Rapa were also affected to a lesser degree than height and lifecycle maturity in all doses; though growth-compensation occurred in the 10 mg/L treatment, inducing changes in survivability and reproductive maturity index ratings. These results suggest a potentially harmful accumulation/sorption dynamic which probably extend to other crops and to animals fed on them. More research is needed to determine the dangers of BPA as both environmental contaminant and consumer toxin.

Introduction

The impact and implications of more than half a century of mass production of polycarbons, plastics, and resins has recently become a hotly-disputed topic within and without the international scientific community. While many plastics have been in mass production for nearly 50 years, only recently have serious objections been raised against the environmental and human costs of such production, described by some as “great environmental concerns.” (Purdom et al., 1994; quoted in Ferrarra et al., 2005).

Industry influence and lobbying is not to be ignored either. In March, 2007, the National Institute of Health dismissed their privately-contracted data analysts (previously in charge of analysis related to BPA and other potentially pathological byproducts of plastics) for conflicts of interest (Sissell, 2007). Industry-led misinformation is also prominent. The website for the “Bisphenol-A Organization” claims that “products made with BPA are safe for their intended uses and pose no known risks to human health” (http://www.bisphenol-a.org/). On the same website there are links to the NIH's National Toxicology Program review of BPA; a 1982 study which found that “the evidence is suggestive of a carcinogenic effect [of BPA] on the hematopoeitic system” (NTP TR-215, 1982): a conclusion which is not mentioned on the website. The NTP report also concluded that “Leukemias in male rats occurred at an incidence that showed a statistically significant positive association with the dose of bisphenol A” (p. 45). A 2001 opinion published by the European Society of Human Reproduction and Embryology stated that “reassessment of models used to test xenobiotics for oestrogenic potency is overdue” (Speraow & Barkley, 2001). A brief review of research linking BPA to animal and plant pathology reveals an extensive corpus of literature: (ECSCF, 2002; EDSTAC, 1998; Staples et al., 1998; Safe, 2000; all quoted in Ferrara, 2006; Ferrara, 2001; B. Scmidt & I. Schupan, 2002; Smith & Taylor, 2007; Stoker et al., 2003; Schirling et al., 2006; MSDS-Sigma-Aldrich, 2006; Atkinson & Roy, 1993, 1995; Hanioka et al., 2000; Nakagawa & Tayama, 2000; Niwa et al., 2000; Colborn et al., 1993; Rice, 2000; Jacobsen & Jacobsen, 1996) to name a few.

According to the Google Trends website, searches for “BPA” and “Bisphenol” have jumped exponentially in 2008, likely due to news that Nalgene Brand water bottles were being recalled from the market due to BPA leaching (sales were banned by law in British Colombia shortly thereafter as well) (http://trends.google.com).

Meanwhile plastics and other polycarbons and resins continue to be mass-produced and marketed in many consumer products ranging from water pipes to babies' bottles, regardless of potential dangers by proven BPA presence (Brede et al., 2003). BPA production in Japan in 1997 was about 400,000 tonnes, and is probably much more than 500,000 tonnes /year worldwide for the last two decades (CMC, 1999; Nakajima, 2004). A German study (Furhacker et al.. 2000) found levels of contamination high enough to pose a danger (especially to aquatic organisms) “emanating” from mostly industrial sites. Especially hazardous were the wastes of paper factories. The study concluded that a significant portion of the BPA was not fully removed through wastewater treatment methods. This study was confirmed in (Hermann et al., 2002).

Environmental BPA contamination is a cause for great anxiety, and so is its presence in consumer products. Many studies have confirmed endocrine disruptor presence in aquatic systems (Dorn et al., 1987; Nasu et al., 2001; Staples, 1998; Cousins et al., 2002; Suzuki et al., 2004; Yamamoto et al., 2001, Ternes et al., 1999). BPA is permittably discharged in some countries (Schmidt & Schupan, 2002; Lobos et al., 1992; Hanioka et al., 2000) and used in all manner of dental sealants and fillings (Ike et al., 2000; Suzuki et al., 2002). Several studies have linked BPA and other hydrophobic plastic by-products to the contamination of the world's oceans to a high degree (Ying & Kookana, 2003; Teuten et al., 2007; Derraik, 2002). Even seals in the pacific northwest have been touched by the contamination (Ross et al., 2004).

According to another article, the presence of other contaminants –heavy metals and metalloid compounds– increased the rate of soil sorption of BPA, indicating what might be termed a 'co-morbidity' of BPA presence in conjunction with other industrial pollutants (Li et al., 2008). Seeming to confirm this idea is (Sun et al., 2008) which concludes that different forms of BPA are absorbed into soils at different rates, with soil organic matter playing a key role in the process.

Understanding the chemical dynamics of sorption of BPA into various soils is the focus of much ongoing current research. Hollrigl-Rosta et al., (2003), discovered that adding additional dissolved organic material did not significantly effect the sorption of BPA in some soils, but did in others. Ying & Kookana (2005) found that aerobic conditions significantly increased the ability of high-concentration organic material soils to degrade BPA and other environmental endocrine disruptors. The authors add that endocrine disruptors (EDC's) should not last long in well-aerated soils, but their persistence “in anaerobic soil may affect soil and groundwater quality and ecosystem.” Kang & Kondo (2002) confirms that degradation of BPA by several strains of bacteria is generally better under aerobic conditions. At least one study questioned the safety of BPA presence in sewage sludge used in agriculture (called biosolids) (La Guardia et al., 2001), and the ability of sewage bacteria to degrade environmental estrogens (Lee & Liu, 2002).

C. Fusca, a microalgae, can remove up to 85% of BPA under ideal photoautotrophic conditions and in certain small concentrations of 40μM and less (Hirooka et al., 2003), but it is unknown if this is possible under more realistic conditions to the same extent.

BPA presence in the rhizospheres of plants seems to cause several different interactions which result in a net phytotoxic effect. First, it probably has adverse effects on the bacteria essential to nitrogen uptake in plants. Many microbes can “detoxify” BPA ( Kang et al., 2006 ) but some cannot. The bacteria Lactococcus Lactis [712] was tested by (Endo et al., 2006) and could remove less than 9% of BPA from media solutions, but could not degrade it without significant additional chemical assistance. This study suggested that “hydrophobic proteins on cell surface may be involved in the BPA-adsorbing ability of lactococci.” Indeed, it is the hydrophobic nature of the chemical xenostrogen compounds that has made them worrisome to researchers. Overall there is some evidence to support a theory of environmental 'co-morbidity' when assessing the hazards of BPA contamination and its phytotoxicity where overall environmental damage may be grossly effected by a multitude of hazards to include EDC's.

Other phytotxic outcomes of BPA sorption in soil have been less well-documented. A handful of studies have demonstrated the negative effects of xenoestrogens on plant cell suspensions and cultures, and fewer still have demonstrated it on plant systems grown hydroponically or in soils (Ferrara et al. 2006). Of these, there have been surprising results. One NATO-funded study compares BPA exposure to gamma ray exposure using Tradescantia micronucleus assays to measure clastogenesis and carcinogenesis, concluding that a relatively small amount of BPA (11.8 mM) has the same cumulative effect as 25 cGy of gamma rays (Kim et al., 2006). This is a strong and telling indictment of the phytotoxicity of BPA. Ferrara et al. (2006) found that BPA exposure did not significantly influence germination or early growth in hydroponically-grown broad-bean, durum wheat, or lettuce plants, but did inhibit tomato root length significantly (p. 3). In their experiment, crops treated with 10 mg/L and 50 mg/L bisphenol A showed many phytotoxic and systemic negative effects, apparent after 21 days. These included morphological alterations of shoots and leaves, soot-blackened roots, leaf chlorosis, and reduced number and size of leaves and roots (ibid). The conclusions drawn by the above study and confirmed by others (Schmidt & Schupan, 2002; Nakajima et al. 2002) seem to indicate that various plant species can absorb large amounts of BPA, and that it should be considered phytotoxic; that EDC's can translocate within the plant, and that “complete biotransformation and detoxification [of BPA] do not occur either in the roots or in the shoots [of some plants]” (Ferrara et al., p. 4).

As the empirical evidence of the phytotoxicity of bisphenol and other chemicals begins to accumulate, it is becoming increasingly concerning (given the ever-increasing contamination of the natural and human environments with that and other harmful chemicals). To prove the toxicological net effect of BPA has been the task for many researchers for the past decade; now the task turns to assessing the full magnitude of the possible environmental crisis, as well as finding means of remediation. Currently, there are few studies which have begun exploring the possibilities of phytoremediation of bisphenol A in the environment. These include Hamada et al., (2002) and Iimura et al., (2007).

The purpose of this paper is to add to the evidence an incomplete analysis of the phytotoxicity of BPA in Brassica Rapa (Wisconsin Fast-start ® mustard). The objective of the experiment performed was to measure only phytotoxic outcomes of BPA contamination through a multi-stage environmental simulation, with the inferred soil sorption, plant uptake, and internal transfer having an end result in visible plant development. By proving the phytotoxicity of EDC's in an easily-reproducible and cost-effective manner, it is hoped that an awareness of the hazardousness of environmental EDC contamination can be improved.

Method & Materials

Plants.

Brassica Rapa (Wisconsin Fast-start ® mustard) seeds were obtained from Carolina Biological Supply, USA. 72 seeds were selected at random from approximately 150 mixed from different containers. All seeds were inspected for uniformity of appearance (size, shape and absence of defects) before selection. Selection and planting occurred at uniform times (within 1 hour for all plants).

Chemical.

Bisphenol A [(2,2-(4,4-dihydroxydiphenol) propane)], 99% purity was obtained from Sigma-Aldrich, USA and dissolved in distilled water at room temperature in concentrations of 10 mg/L and 50 mg/L. A total of 1.50 liters of each solution was mixed, and allowed to stand for approximately 1 day in sealed containers before irrigation. Distilled water was used for the controlled variable plants.

Soil.

Potting soil mix was obtained from WCC lab stock of a mixture of commercially-available brands. Soil was inspected for uniformity of appearance and consistency before planting. Soil was mixed and moistened with distilled water before selection, and randomly selected for plantings. Amount of soil was standardized at approximately 1.5 g.

Watering regimes and housing.

Watering regimes were constructed to form a reservoir (made of sealed tupperware®-brand container) and a cloth wicking system to allow for continuous irrigation. Styrofoam® chambered planters (also obtained from Caolina Biological Supply, USA) with four separate chambers for soil (with holes and additional wicks at the bottom for irrigation) were placed on top of the regimes. All materials were selected at random from lab stocks, and thoroughly inspected for uniformity and pre-cleaned before usage.

Lighting and temperature.

Full-spectrum lighting was set up in a lab chamber to allow for thorough lighting and uniformity of light brightness and quality. Lighting was set on a daily timer for 12 continuous hours of light followed by 12 hours of dark. Light was provided during planting as well. All measurements occurred during lighted hours to mitigate external effects, and lighting followed a general daylight schedule to allow for any other seasonal or diurnal cues. Temperature remained at constant room temperature for the duration of the experiment.

Planting, depth, initial irrigation.

Planting of seeds was standardized in soil at 5-7 mm depth, and within an hour for all seeds. Initial irrigation of treatment or control water was given via dropper until all wicks and soil were thoroughly irrigated. Two repetitions of three regimes for each variable (six total planters for 10 mg/L, 50 mg/L and 0.0 mg/L, each with four seeds in separate chambers) were planted. Seeds were given separate chambers to mitigate any root or placement competition, and irrigation was checked hourly for the first 4 hours to ensure thorough irrigation. All plants within each chamber were therefor of the same treatment.

Placement.

Placement within the lighting chamber was randomized after the first day on a semi-daily basis to minimize any potential effects to plant growth. Watering regimes within treatment groups were randomly switched twice during the experiment to mitigate any potential “block effect” on plant growth.

Continuing irrigation.

During the course of the 26 days, plants were continually irrigated through the watering regimes and checked on a daily basis for thoroughness of irrigation. Plants were hand-irrigated on a standardized basis directly following every measurement session to ensure continued uninterrupted growth.

Measurements.

Measurements were taken at standard times at 4, 8, 10, 15, 19, 23, and 26 days. Standardization of practices included measurement tools and techniques, as well as timing and thoroughness. Variables measured for analysis were turgidity/flaccidity (subjective yes/no criteria), number of buds (objective criteria), number of flowers( objective criteria), number of leave-halves (objective criteria), presence of obvious deformities (subjective yes/no criteria), and cumulative height above soil top: accurate to 1mm (objective criteria). Height was measured using standardized instrument (ruler) by reading the tallest portion of the plant. Leave-halves were counted in order to reduce the margin of error due to presence of deformities of leaves. Turgidity and flaccidity were subjective criteria determined on the basis of deviation from documented deformities/growth-patterns but not included in the final statistical analysis of the results.

Analysis.

Analysis was performed using simplistic variable-analysis in an automated spreadsheet format. See Results below.

Results & Discussion

Definition of terminology.

Variations in sexual maturity were measured primarily by determination of number of buds and number of flowers. An index of these co-variables was created, termed here “Sexual Maturity Rating,” with the mathematical formula of M = (1/3) Tbuds + Tflowers where T(buds) is total number of buds and T (flowers) is total number of flowers where sexual maturity reached critical stages on a variable-criteria timetable (budding by day 15, flowering by day 23).

A vivacity index was determined in order to combine multiple variables for single analysis. The formula for this is:

formula

where H is cumulative height in mm, Td is time in days, Nlh is number of leave halves, MS is the sexual maturity rating described above, and Nd is the number, if any, of defects on the plant. Having these multi-variable ratings allowed for a more general analysis of phytotoxic effects of BPA on the plants.

Results and discussion.

Generally, the phytotoxic effects of BPA were apparent at 50 mg/L levels in all measured variables, but they were also apparent in the sexual maturity ratings and late growth curves for the 10mg/L treatment. The overall germination of Brassica Rapa was not significantly inhibited by BPA presence but early growth differentiation began emerging by day 10. By day 19 there is a significant difference in height, producing a growth-equilibrium shift that lasted through the rest of the experiment (see figures 1, 2.)

figure 1
(Figure 1.)

figure 2
(Figure 2.)

Only one negative value appeared in growth charting: control treatments on the last measurement. This apparent “shrinkage” in the non-BPA treated group is actually the result of the weight of buds and flowers bending the plants slightly downward as well as a halt in the growth patterns of the plants. Whereas the sexual maturity of the control group was greater at this point in the experiment (see figures 3 & 4) and the growth of the BPA-treated plants was somewhat retarded, it can be said with some assurance that the shrinkage in the control group was quite a natural occurrence.

Interestingly, the growth of the treated plants did not stop as did the control, though the 10mg/L treatment plants did slow significantly between days 23 and 26. This may be due to the BPA inhibition of cell-elongation and / or division (Vaughan & Ord, 1990: as quoted in Ferrara et al., 2006) because generally the plants' sexual maturity stages still occurred in marked stages transcending treatment variable. These findings seem to corroborate those of Ferrara et al. (2006) in the assessment of the effects of BPA during germination.

Sexual maturity ratings (see definition above) demonstrate more contiguous changes by addition of BPA. Because of the time-specific categorization of the sexual maturity ratings (i.e. flowering or budding by a set number of days) the differences between the 10mg/L treatment and control plants becoming more acute, while the obvious interference of BPA on the reproductive maturation at the 50mg/L level is rather obvious.

figure 3
(Figure 3.)

Because the number of flowers reaching maturity is a function of the number of buds, and because of the fact that not all buds reached maturity to become flowers, there is a slight drop sexual maturity curves between days 15 and 19. However, most plants had flowered by day 23 in all variables, and the disparity in ratings can be clearly correlated to BPA presence by that time.

figure 4 
(Figure 4.)

The overall vivacity rating (VR) was a rating combining most of the dependent variables in such a way as to determine systemic effects of BPA presence in plants. The results of that are as shown in figures 5-6.

figure 5
(Figure 5.)

figure 6
(Figure 6.)

Overall, more visible deformities occurred in BPA-treated plants than in control plants, but the same level of correlation between treatments that was evident in the sexual maturity ratings is not apparent in the vivacity rating. (Although the net phytotoxic effect of the 50mg/L treatment is still evident). This change is most likely due to compensation of growth rates as a function of BPA presence.

The reasons for the demonstrated overall net phytotoxicity are probably multi causal with regards to bisphenol A presence. While full analysis was avoided in this study, it is suggested by Ferrara et al. (2006) and Schmidt & Schupan (2002) that there are various aspects to BPA toxicity, possibly including the release of acidic root exudates, and changes in the rhizospheres including microbial and electrolyte concentration changes. Further analysis is needed to determine specific pathology of all xenoestrogens in plants.

Conclusions

While the exact pathology of Bisphenol A disruption to plant organisms is still somewhat unknown, the evidence of its phytotoxicity is relatively abundant. By empirically measuring the systemic effect of BPA in a simulated natural environment, we can conclude that the results of previous research in hydroponically-grown plants and other lab settings holds true in more natural ones as well. There is strong evidence that BPA contamination of the environment plays a 'co-morbid' role along with other hazardous contaminants; there is also strong evidence for a theory of EDC bioaccumulation in crops and non-crop plants. More importantly, as the specific science of the phytoremediation, degradation, and complete detoxification of BPA and its metabolites remains unknown, it posses a possible threat to human and animal consumption through the food chain. More research is needed in order to determine the true extent of this threat.

It is hoped that this experiment can be used both as a demonstration of the systemic effects of the environmental contamination of BPA, and as a simple way to prove the general phytotoxicity of synthetic estrogens and other EDC's produced by man.

The author would like to thank John Rousseau, the WCC lab staff, Doreen Dewell, Beth Tyne, Steve DeRoy, Dryw Hopps and Marie Marchand.

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