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Archive for the ‘Safe Pest Control’ Category

http://advances.sciencemag.org/content/4/7/eaas9024
Biodegradation of synthetic polymers in soils: Tracking carbon into CO2 and microbial biomass

Abstract

Plastic materials are widely used in agricultural applications to achieve food security for the growing world population. The use of biodegradable instead of nonbiodegradable polymers in single-use agricultural applications, including plastic mulching, promises to reduce plastic accumulation in the environment. We present a novel approach that allows tracking of carbon from biodegradable polymers into CO2 and microbial biomass. The approach is based on 13C-labeled polymers and on isotope-specific analytical methods, including nanoscale secondary ion mass spectrometry (NanoSIMS). Our results unequivocally demonstrate the biodegradability of poly(butylene adipate-co-terephthalate) (PBAT), an important polyester used in agriculture, in soil. Carbon from each monomer unit of PBAT was used by soil microorganisms, including filamentous fungi, to gain energy and to form biomass. This work advances both our conceptual understanding of polymer biodegradation and the methodological capabilities to assess this process in natural and engineered environments.

INTRODUCTION

Modern agriculture heavily relies on the use of plastic materials in various applications, a practice coined plasticulture. Mulching with plastic films is a major application with a global market volume of approximately 2 × 106 tons per year (1). Mulch films are placed onto agricultural soils to improve conditions for plant growth while lowering consumption of water, herbicides, and fertilizer and also minimizing soil erosion (1, 2). Because of these benefits, mulching with plastic films helps to secure food for the growing world population. However, mulch films are commonly composed of nonbiodegradable polyethylene and, therefore, accumulate in agricultural soils and surrounding receiving environments if incompletely retrieved after use. These accumulations have negative ecologic and economic impacts, including decreased soil productivity (35). A promising strategy to overcome these risks is to use mulch films composed of polymers that biodegrade in soils (1, 68).

Biodegradation of polymers requires microorganisms to metabolize all organic components of the polymer. Biodegradation in soil involves several key steps (Fig. 1): (i) colonization of the polymer surface by microorganisms, (ii) secretion of extracellular microbial enzymes that depolymerize the polymer into low–molecular weight compounds, and (iii) microbial uptake and utilization of these compounds, incorporating polymer carbon into biomass or releasing it as CO2 (9).

Fig. 1 Key steps in the biodegradation of polymers in soils.

Microorganisms colonize the polymer surface and secrete extracellular enzymes that depolymerize the polymer. The formed low–molecular weight hydrolysis products are taken up by the microorganisms and used both for energy production, resulting in the formation of CO2, and for the synthesis of cellular structures and macromolecules, resulting in incorporation of polymer-derived carbon into the microbial biomass. The boxes on the right depict the analytical methods we used to study these steps. NMR, nuclear magnetic resonance.

Fig. 1 Key steps in the biodegradation of polymers in soils.Microorganisms colonize the polymer surface and secrete extracellular enzymes that depolymerize the polymer. The formed low–molecular weight hydrolysis products are taken up by the microorganisms and used both for energy production, resulting in the formation of CO2, and for the synthesis of cellular structures and macromolecules, resulting in incorporation of polymer-derived carbon into the microbial biomass. The boxes on the right depict the analytical methods we used to study these steps. NMR, nuclear magnetic resonance.Here, we examined each of the above steps for poly(butylene adipate-co-terephthalate) (PBAT), an aliphatic-aromatic statistical copolyester of large importance in the market of biodegradable mulch films (7). While previous studies provided indirect indications for PBAT biodegradation in soils based on determining PBAT mass loss and changes in its physicochemical properties (1012), we here use a novel workflow using stable carbon isotope-labeled PBAT to directly and unequivocally demonstrate its biodegradation in soil (table S1). This workflow included incubation of 13C-labeled polymer films in soil with continuous quantification of polymer-derived 13CO2 by isotope-specific cavity ring-down spectroscopy (CRDS) (13). The use of 13C-labeled polymers allowed us to distinguish polymer-derived CO2 from CO2formed by soil organic matter mineralization. After incubation, we imaged the polymer film surfaces using scanning electron microscopy (SEM) and demonstrated the incorporation of polymer-derived 13C into the biomass of film-colonizing microorganisms using element-specific, isotope-selective nanoscale secondary ion mass spectrometry (NanoSIMS) (14). We studied three PBAT variants that had similar physicochemical properties and comparable total 13C contents, but varied in the monomer that contained the 13C-label [that is, butanediol (P*BAT), adipate (PB*AT), or terephthalate (PBA*T)] (Fig. 2A and table S2). The use of these variants allowed us to follow biodegradation of all PBAT building blocks. The presented workflow is a novel approach to study the fundamental steps in polymer biodegradation in complex systems (1517).

DISCUSSION

This work presents an experimental approach to study polymer biodegradation in soils and to assess the key steps involved in this process: microbial polymer colonization, enzymatic depolymerization on the polymer surface, and microbial uptake and utilization of the released low–molecular weight compounds. Central to the approach is the use of polymer variants that are 13C-labeled in all monomer units of the polymer, thereby allowing us to assess whether all organic components of the polymer material are used by soil microorganisms. The label further allows tracing of polymer-derived carbon into both CO2 and microbial biomass. Using this approach, we demonstrate here the biodegradability of PBAT in soil. Biodegradability renders PBAT a more environmentally friendly alternative to persistent polymer materials for use in plasticulture, including single-use applications such as plastic mulching. Our results further imply that incorporation of polymer-derived carbon into microbial biomass needs to be taken into consideration in regulatory guidelines for determining biodegradability of polymers. Currently, these guidelines are solely based on extents of CO2 formation. Furthermore, the finding of subcellular structures within PBAT-colonizing fungi highly enriched in polymer-derived carbon might represent compartments in which carbon is stored (for example, in the form of neutral lipids) when fungi are limited by the availability of nutrients other than carbon (22). These limitations are plausible for microorganisms growing on PBAT and other polymers that do not contain nitrogen and phosphorous. If these limitations occur, increasing the availability of soil nutrients to microorganisms colonizing the polymer surface is expected to enhance polymer biodegradation.

This work demonstrates PBAT biodegradation in a selected agricultural soil over 6 weeks of incubation. Future studies extending on this work will need to assess variations in the rates and extents of PBAT mineralization among different agricultural soils, also over longer-time incubations. Furthermore, we propose studies that are directed toward identifying soil microorganisms that are actively involved in PBAT biodegradation. While the NanoSIMS-based approach presented here allows us to unambiguously demonstrate incorporation of polyester carbon into soil microbial biomass, it is not a high-throughput technique. Alternative approaches, including the extraction of targeted biomolecules from soils containing 13C-labeled polymers followed by quantifying the 13C contents in the extracted molecules, will allow us to analyze larger sample sets and thereby to systematically determine potential variations among soil microorganisms in the extent to which they incorporate polymer-derived carbon into their biomass.

MATERIALS AND METHODS

Experimental design

The objective of this study was to develop an experimental approach to demonstrate biodegradation of PBAT in an agricultural soil. As biodegradation includes mineralization of PBAT carbon to CO2, as well as the incorporation of PBAT-derived carbon into the biomass of soil microorganisms, we addressed both of these processes in controlled laboratory experiments. We followed PBAT mineralization during soil incubation using an isotope-specific CRDS for the quantification of formed CO2. For each of the three PBAT variants, we simultaneously incubated seven films in one incubation bottle filled with soil to allow precise quantification of PBAT mineralization to CO2. The soil incubations were terminated after 6 weeks (that is, when approximately 10% of the PBAT carbon had been mineralized) to ensure that PBAT films were still intact for the subsequent imaging analyses. We revealed incorporation of PBAT-derived carbon into biomass using NanoSIMS, which enabled identification of subcellular features and determination of the carbon isotope composition of the PBAT film surface and the colonizing microorganisms at submicrometer spatial resolution. The low throughput of this high-end topochemical analysis technique constrained the number of collected images for soil-incubated films to two images for each of the three PBAT variants including replicate films. We note that we did not exclude any data or outliers from our analysis.

Polyesters, monomers, soil, and enzymes

Polyesters were provided by BASF SE and synthesized as previously described (23, 24). The physicochemical properties of the polyesters are listed in table S2. To obtain similar 13C contents for the three PBAT variants (that is, PB*AT, P*BAT, and PBA*T), synthesis of all variants was performed with defined ratios of labeled to unlabeled monomers. The three PBAT variants were free of chemical additives.

The 13C-labeled monomers 1,6-13C2-adipate and 13C4-butanediol used for PBAT synthesis and for soil incubation studies were purchased from Sigma, with more than 99% of the indicated positions in the monomer containing 13C. We obtained 1-13C1-terephthalate from dimethyl 1-13C-terephthalate purchased from Sigma. To obtain the free diacid, we dissolved dimethyl 1-13C-terephthalate in 2:1 water/tetrahydrofuran (5 mg in 2.4 ml), added 25 μl of a sodium hydroxide solution [37% (w/w)], and stirred the solution at room temperature for 2 hours. The solvent was then carefully removed under reduced pressure to obtain the hydrolysis product 1-13C1-terephthalate (confirmed by 1H NMR).

For PBAT and monomer incubations in soils under controlled laboratory conditions, we used agricultural soils from the agricultural center Limburgerhof (Rhineland-Palatinate, Germany). Physicochemical properties of the soils are provided in table S1. The soils were air-dried to a humidity of 12% of the maximum water-holding capacity of the soil, passed through a 2-mm sieve, and stored in the dark at 4°C for 12 months before use in the incubation experiments.

R. oryzae lipase was purchased as a powder from Sigma (catalog no. 80612). FsC was obtained as a solution from ChiralVision B.V. (Novozym 51032). Stock solutions of both enzymes in water were stored at −20°C.

Preparation of PBAT films and soils for incubation experiments

We prepared two sets of solvent-cast PBAT films that differed in the way that the PBAT films were attached to the silicon wafer substrates. For the first set, we solvent-cast PBAT films by adding three times 15 μl of a PBAT solution in chloroform [concentration, 5% (w/w)] onto a square-cut antimony-doped silicon wafer platelet (7.1 mm × 7.1 mm × 0.75 mm, Active Business Company). In between the additions of the polymer solutions, we allow the chloroform to evaporate. This procedure resulted in a PBAT mass of approximately 3 mg per wafer. Before incubation in soil, the solvent-cast polyester films were stored in the dark at room temperature for 1 week to ensure complete evaporation of the solvent (chloroform). PBAT variants from this first set were used for PBAT mineralization experiments (Fig. 2B), SEM imaging (Fig. 2C), and NanoSIMS imaging (Figs. 3 and 4 and fig. S8).

For the second set of PBAT films, we pretreated the silicon wafer platelets with Vectabond (Vector Laboratories, catalog no. SP-1800) before solvent casting of the polyester films. This second set of PBAT films was included to test whether the adhesion of the PBAT to the Si surface can be improved by this modified protocol. For the pretreatment, we exposed the wafers to a 1:50 diluted solution of Vectabond in acetone, subsequently dipped them into MilliQ water (Barnstead Nanopure Diamond), and dried them in a stream of N2. PBAT variants from this set were used only to determine PBAT mineralization (fig. S1), but not for SEM and NanoSIMS imaging.

We prepared the soil for PBAT incubations by adding MilliQ water to the soil to adjust its water content to 47% of its maximum water-holding capacity. We subsequently transferred 60 g of the soil into each of the incubation vessels (100-ml glass Schott bottles). We prepared a total of nine incubation bottles in three sets of three bottles (see below). The soils were then preincubated at 25°C in the dark for 1 week.

After soil preincubation, we transferred the wafers carrying the solvent-cast polyester films into the soils in the incubation bottles. We added seven wafers to each incubation bottle. The wafers were spaced apart by at least 1 cm. All wafers were positioned upright in the soil. The three bottles of the first set each contained films of one of the three differently labeled PBAT variants obtained by direct solvent casting. The three bottles of the second set were identical to the first set except for the wafers, which were pretreated with Vectabond before solvent casting. The three bottles in the third set served as controls and contained soil but no PBAT films. All bottles were incubated for 6 weeks at 25°C in the dark. We note that our study therefore does not address potential effects of ultraviolet irradiation–induced changes in the structure of PBAT on its biodegradability. Over the course of the incubation, we gravimetrically determined the water content of the soils at defined time intervals. To sustain a constant soil water content, amounts of water that were lost from the soil through evaporation were replenished by adding corresponding amounts of MilliQ water.

Preparation and SEM imaging of soil-incubated PBAT films

After 6 weeks of incubation in soil, we carefully removed the silicon wafers carrying the PBAT films from the soils. To chemically fix the cells attached to the surfaces of the PBAT films, we directly transferred the films into a freshly prepared fixation solution (pH 7.4) containing glutaraldehyde (2.5%), sodium cacodylate (0.1 M), sodium chloride (0.1 M), potassium chloride (3 mM), and sodium phosphate (0.1 M). The films were exposed to this solution for 20 min at 25°C and subsequently transferred to a solution of OsO4 in MilliQ water (1%) for 30 min of exposure on ice. Finally, we dehydrated the films in a series of water/ethanol solutions of increasing concentrations (70%, 5 min; 95%, 15 min; 100%, 2 × 20 min), followed by critical point drying of the samples with liquid CO2 (Baltec CPD 030). Critical point drying resulted in detachment of the PBAT films from the wafer. To reattach the films to the wafers for further analyses, we used a double-sided adhesive, electrically conducting carbon tape (Ted Pella, product no. 16084-1). Directly after mounting the films onto the wafers with carbon tape, thin films of platinum (thickness, 10 nm) were deposited on the samples using a sputter coater (Baltec SCD 500). SEM was conducted on a Zeiss Supra 50 VP. Imaging was performed with a secondary electron detector at a working distance of 4.0 mm and an electron high tension of 5.0 kV. These films were also used for NanoSIMS analysis (see below).

PBAT films from the second set, for which wafers were pretreated with Vectabond before solvent casting of PBAT (see above), also detached from the wafers. We decided to reject further analysis of these films (that is, SEM and NanoSIMS).

PBAT film imaging by NanoSIMS

NanoSIMS measurements were performed on a NanoSIMS NS50L (Cameca) at the Large-Instrument Facility for Advanced Isotope Research (University of Vienna). Before data acquisition, analysis areas were presputtered by scanning of a high-intensity, slightly defocused Cs+ ion beam (beam current, 400 pA; spot size, approximately 2 μm). To avoid crater edge effects, scanning during presputtering was conducted over square-sized areas with an edge length exceeding the frame size of the subsequently recorded images by at least 15 μm. Every data set acquired on the soil-incubated polymer films contains image data recorded from (at least) two distinct depth levels, accessed by sequential presputtering with Cs+ ion fluences of 5.0 × 1016 and 2.0 × 1017 ions/cm2, respectively. Application of the lower ion dose density enabled sampling of all cells within the analysis areas, irrespective of their size and/or morphology, whereas the extended presputtering allowed us to gain insight into cellular features contained within the lumen of bulky cells such as fungal hyphae (see, for example, Fig. 4).

Imaging was conducted by sequential scanning of a finely focused Cs+ primary ion beam (2-pA beam current) over areas ranging from 45 × 45 μm2 to 70 × 70 μm2 at a physical resolution of approximately 70 nm (that is, probe size) and an image resolution of 512 × 512 pixels. If not stated otherwise, images were acquired as multilayer stacks with a per-pixel dwell time of 1.5 ms per cycle. 12C, 13C, 12C12C, 12C13C, 12C14N, 31P, and 32S secondary ions as well as secondary electrons were simultaneously detected, and the mass spectrometer was tuned for achieving a mass resolving power of >9.000 (according to Cameca’s definition) for detection of C2 and CN secondary ions. Image data were analyzed with the ImageJ plugin OpenMIMS, developed by the Center for NanoImaging (27). Secondary ion signal intensities were corrected for detector dead time (44 ns) and quasi-simultaneous arrival (QSA) of secondary ions. Both corrections were performed on a per-pixel basis. QSA sensitivity factors (“beta values”) were obtained from measurements on dried yeast cells, yielding 1.1, 1.06, and 1.05 for C, C2, and CN secondary ions, respectively. Before stack accumulation, images were corrected for positional variations originating from primary ion beam and/or sample stage drift. ROIs were manually defined on the basis of 12C14N secondary ion signal intensity distribution images and cross-checked by the topographical/morphological appearance indicated in the simultaneously recorded secondary electron images (see fig. S10). While each cell from unicellular organisms was assigned to an individual ROI, image regions within the polyester surfaces and hyphae were segmented into multiple ROIs. Throughout the article, the carbon isotope composition is displayed as the 13C/(12C + 13C) isotope fraction, given in at%, calculated from the C and C2 secondary ion signal intensities via 13C/(12C + 13C) and 13C12C/(2⋅12C12C + 13C12C), respectively. Owing to superior counting statistics, all carbon isotope composition data shown in the article were inferred from C2signal intensities. We note that we did not observe any significant differences between 13C content values inferred from C2 signal intensities versus C signal intensities. For the line scan analyses displayed in Fig. 4, C2 normalized C14N signal intensities were used as an indicator of the relative nitrogen content {calculated via [12C14N (1 + 13C/12C)]/[12C13C + 12C2 (1 + (13C/12C)2)], whereby the term 13C/12C refers to the 13C-to-12C isotope ratio, calculated from the C2 signal intensities via 13C12C/(2⋅12C12C)}. This quantity formally refers to the relative nitrogen-to-carbon elemental ratio and was used in favor of the relative nitrogen concentration, which is inferable from C normalized C14N signal intensities, to minimize artifacts arising from the considerable topography within the areas of the fungal hyphae (28).

SUPPLEMENTARY MATERIALS

Supplementary material for this article is available at http://advances.sciencemag.org/cgi/content/full/4/7/eaas9024/DC1

Supplementary Materials and Methods

Fig. S1. Mineralization of PBAT films.

Fig. S2. NMR analysis of enzymatic hydrolysis products of PBAT films I.

Fig. S3. NMR spectra of terephthalate, adipate, and butanediol.

Fig. S4. NMR analysis of enzymatic hydrolysis products of PBAT films II.

Fig. S5. NMR analysis of enzymatic hydrolysis products of PBAT films III.

Fig. S6. NMR analysis of enzymatic hydrolysis products of PBAT films IV.

Fig. S7. Mineralization of terephthalate, adipate, and butanediol.

Fig. S8. NanoSIMS analysis of PBAT films after soil incubation I.

Fig. S9. Control experiment for NanoSIMS analysis I.

Fig. S10. Definition of ROIs.

Fig. S11. Control experiment for NanoSIMS analysis II.

Fig. S12. NanoSIMS analysis of PBAT films after soil incubation II.

Table S1. Soil characterization.

Table S2. Characterization of PBAT variants.

Supplementary Appendix. Calculations of the carryover during NanoSIMS measurements.

References (2933)

This is an open-access article distributed under the terms of the Creative Commons Attribution-NonCommercial license, which permits use, distribution, and reproduction in any medium, so long as the resultant use is not for commercial advantage and provided the original work is properly cited.

Acknowledgments: We thank S. Probst, M. Jaggi, and F. Strasser for their help with growing E. coli, performing IRMS measurements, and NanoSIMS control sample preparation and data analysis, respectively. Funding: M.T.Z., T.F.N., R.B., H.-P.E.K., K.M., and M.S. thank the Joint Research Network on Advanced Materials and Systems of BASF SE and ETH Zürich for scientific and financial support. M.W. and A.S. were supported by the European Research Council Advanced Grant project NITRICARE 294343. D.W. was supported by the European Research Council Starting Grant project DormantMicrobes 636928. SEM imaging was performed at the Center for Microscopy, University of Zurich. Author contributions: M.T.Z., A.S., D.W., H.-P.E.K., K.M., and M.S. designed the study. M.T.Z., A.S., T.F.N., and R.B. performed experiments. All authors contributed to the writing of the manuscript. Competing interests: The authors declare that they have no competing interests. Data and materials availability: All data needed to evaluate our conclusions are present in the paper and/or the Supplementary Materials. Additional data related to this paper may be requested from the authors.

  • Copyright © 2018 The Authors, some rights reserved; exclusive licensee American Association for the Advancement of Science. No claim to original U.S. Government Works. Distributed under a Creative Commons Attribution NonCommercial License 4.0 (CC BY-NC).

 

 

 

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Household Chemical Exposures and the Risk of Canine Malignant Lymphoma, a Model for Human Non-Hodgkin’s Lymphoma

Biki B. Takashima-Uebelhoer,1 Lisa G. Barber,2 Sofija E. Zagarins,3 Elizabeth Procter-Gray,4 Audra L. Gollenberg,5Antony S. Moore,2,6 and Elizabeth R. Bertone-Johnson1,2

https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3267855/

Epidemiologic studies of companion animals offer an important opportunity to identify risk factors for cancers in animals and humans. Canine malignant lymphoma (CML) has been established as a model for non-Hodgkin’s lymphoma (NHL). Previous studies have suggested that exposure to environmental chemicals may relate to development of CML..

Conclusions. In summary, findings of this study suggest that exposure to certain types of lawn care chemicals may increase the risk of malignant lymphoma in dogs. Additional studies are needed to further evaluate the effects of specific chemical components of lawn care products on risk of canine malignant lymphoma, and may potentially contribute to human NHL as well.

Results suggest that use of some lawn care chemicals may increase the risk of CML. Additional analyses are needed to evaluate whether specific chemicals in these products may be related to risk of CML, and perhaps to human NHL as well.

Keywords: Lymphoma, Non-Hodgkin, Dogs, Epidemiology, Case-Control Studies, Specialty Uses of Chemicals

(more…)

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Household Chemical Exposures and the Risk of Canine Malignant Lymphoma, a Model for Human Non-Hodgkin’s Lymphoma

In summary, findings of this study suggest that exposure to certain types of lawn care chemicals may increase the risk of malignant lymphoma in dogs. Additional studies are needed to further evaluate the effects of specific chemical components of lawn care products on risk of canine malignant lymphoma, and may potentially contribute to human NHL as well.

Lawn Chemicals Linked to 2 Types of Cancer in Dogs

According to a study conducted over a 6 year period at the Foster Hospital for Small Animals at Tufts University’s Cummings School of Veterinary Medicine, exposure to lawn pesticides, specifically those applied by professional lawn care companies, raised the risk of canine malignant lymphoma (CML) by as much as 70%

Epidemiologic studies of companion animals offer an important opportunity to identify risk factors for cancers in animals and humans. Canine malignant lymphoma (CML) has been established as a model for non-Hodgkin’s lymphoma (NHL). Previous studies have suggested that exposure to environmental chemicals may relate to development of CML.

Methods We assessed the relation of exposure to flea and tick control products and lawn-care products and risk of CML in a case-control study of dogs presented to a tertiary-care veterinary hospital (2000–2006). Cases were 263 dogs with biopsy-confirmed CML. Controls included 240 dogs with benign tumors and 230 dogs undergoing surgeries unrelated to cancer. Dog owners completed a 10-page questionnaire measuring demographic, environmental, and medical factors.

Results After adjustment for age, weight, and other factors, use of specific lawn care products was associated with greater risk of CML. Specifically, the use of professionally applied pesticides was associated with a significant 70% higher risk of CML (odds ratio(OR)=1.7; 95% confidence interval (CI)=1.1–2.7). Risk was also higher in those reporting use of self-applied insect growth regulators (OR = 2.7; 95% CI=1.1–6.8). The use of flea and tick control products was unrelated to risk of CML.

Conclusions Results suggest that use of some lawn care chemicals may increase the risk of CML. Additional analyses are needed to evaluate whether specific chemicals in these products may be related to risk of CML, and perhaps to human NHL as well. (more…)

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The developing world is awash in pesticides. By Aleszu Bajak

http://www.vox.com/2016/7/3/12085368/developing-world-pesticides

In today’s globalized world, it is not inconceivable that one might drink coffee from Colombia in the morning, munch cashews from Vietnam for lunch, and gobble grains from Ethiopia for dinner. That we can enjoy these products is thanks, in large part, to expanded pesticide use across the developing world.

Every year, some 3.5 billion kilograms (7.7 billion pounds) of pesticides — a catchall term for the herbicides, insecticides, and fungicides applied to crops from seed to harvest — are used to preserve the quality and quantity of fruits, vegetables, and grains. Herbicides, such as Monsanto’s weed killer glyphosate, make up the bulk of the pesticides applied worldwide. 

In the developing world, where swelling populations, increased urbanization, and growing economies create a demand for ever more food — produced quickly and inexpensively — pesticide application rates are rising. Bangladesh and Thailand have quadrupled their pesticide use since the early 1990s, while Ghana, Ethiopia, and Burkina Faso, countries newer to the pesticide game, have seen a tenfold increase over the same period, according to the Food and Agriculture Organization of the United Nations. (more…)

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The Myths of Safe Pesticides by André Leu.  Andre Leu challenges conventional farming methods by refuting the myths that surround the use and understanding of pesticides.

In his new book The Myths of Safe Pesticides, Andre Leu  exposes the dangers of these chemicals. The chemical-based conventional agriculture industry claims that the synthesized concoctions they sell as pesticides, herbicides, and insecticides are safe when used as directed, but does the scientific evidence truly support their assertions?

Lecturer André Leu delves into a wealth of respected scientific journals to present the peer-reviewed evidence that proves the claims of chemical companies and pesticide regulators are not all they seem.
André Leu challenges conventional farming methods by refuting the myths that surround the use and understanding of pesticides. He exposes the dangers of these chemicals. http://www.acresusa.com/the-myths-of-safe-pesticides

(more…)

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Why the Precautionary Principle is so important to the health of our families- especially our children and grandchildren
If you opted to NOT spray  PESTICIDES at your home or school you made the wise decision  
This wise decision is the Precautionary Principle. 
And It is such a very fortunate decision because on the 20th of March 2015, the International Agency for Research on Cancer (IARC), the specialized cancer agency of the World Health Organization, assessed the carcinogenicity of pesticides, and classified the herbicide Glyphosate and the insecticides Malathion and Diazinon as probable carcinogenic to humans. 

(more…)

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Neurodevelopmental Disorders and Prenatal Residential Proximity to Agricultural Pesticides: The CHARGE Study.

This study of ASD strengthens the evidence linking neurodevelopmental disorders with gestational pesticide exposures, particularly organophosphates, and provides novel results of ASD and DD associations with, respectively, pyrethroids and carbamates

Children of mothers who live near agricultural areas, or who are otherwise exposed to organophosphate, pyrethroid, or carbamate pesticides during gestation may be at increased risk for neurodevelopmental disorders.

Janie F. Shelton,1 Estella M. Geraghty,2 Daniel J. Tancredi,3,4 Lora D. Delwiche,1 Rebecca J. Schmidt,1 Beate Ritz,5,6,7 Robin L. Hansen,3,8 and Irva Hertz-Picciotto1,8

http://ehp.niehs.nih.gov/1307044/ (more…)

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