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CHAPTER 6. METHOD FOR MEASURING HELMINTH EGG CONCENTRATIONS IN WASTEWATER 6.1. INTRODUCTION This chapter presents the results completed for the objective of determining the most appropriate method for monitoring environmental samples for parasitic helminth eggs in Mexicali research laboratories – same comment as other chapters – don’t try to paraphrase the objective – list is exactly how it is stated in the first chapter. The literature citing the available methods for detection of helminth eggs in wastewater and biosolids was first reviewed to find a simple, fast, and economical method with good recoverability. Once the method was chosen, training in Dr. Dwight Bowman’s parasitology laboratory at Cornell University ensured proper sample processing and proper identification of parasitic helminth eggs. This method was further practiced during the spring, summer, and fall sampling trips, and details of the transfer to Mexicali research laboratories are given. Finally, the illustrated step-by-step protocol is presented in both English and Spanish. 6.1.1. Importance of monitoring for intestinal parasites In developing countries, infections due to helminthes are endemic in the population, and the risk to public health should be analyzed when raw or treated wastewater is used for agricultural purposes. Approximately 4.5 billion people worldwide are infected with parasitic helminthes, and epidemiological studies have correlated infection with the reuse of raw wastewater (WHO, 1989, Cifuentes et al., 1993). The World Health Organization publishes standards for the reuse of wastewater in agricultural, limiting wastewater to 1 (viable) helminth egg/ liter of water when used during irrigation (WHO, 1989). Monitoring for fecal coliforms and E. coli is a common practice for indication of pathogenic bacteria, but routine monitoring for pathogenic helminth eggs is difficult due to lack of laboratory personnel experienced in identifying helminth eggs (Westcot, 1997). Currently, very little data for helminth concentrations in raw wastewater and polluted surface waters exists. Mexico has adopted the WHO helminth egg guideline of 1 egg/l, but does not require viability testing. Public works agencies, including CESPM would like to perform routine monitoring for helminth eggs on reuse wastewaters, but are hesitant due to the lack of detection experience. 6.1.2. Comparison and selection of helminth egg detection method

The available detection techniques for Ascaris lumbricoides eggs, generally consist of two parts—separating the eggs from the environmental matrix through various sedimentation or flotation steps, and enumeration and viability determinations using direct microscopy (Ayres et al., 1996, Nelson et al., 2001). Numerous methods, including those recommended by WHO and the USEPA, have been published for identifying and enumerating helminthes, each having their own advantages and disadvantages. These methods were originally developed for use in medical parasitology for feces samples, and are not suitable for the low concentrations of eggs in wastewater samples. Because testing has only recently gained popularity, there is a lack of quality assurance and quality control (QA/QC) data for most methods (Bowman et al., 2003). Many methods are time consuming, and do not have a high percentage of recoverability (< 50%),

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and some use hazardous reagents like ether, not appropriate for laboratories with limited protective equipment (Ayres et al., 2001).

The recommended WHO ‘Bailenger’ method, adapted from fecal analyses for environmental samples is inexpensive, quick, and has the ability to recover a wide range of helminth species, but the percent recovery of eggs is not known. This method was developed for use in laboratories internationally in an effort to standardize the method for helminth egg detection, and an illustrated step-by-step protocol is available on the internet. Disadvantages to this method are due to its use of McMaster slides to examine wastewater samples during microscopy. These slides are only recommended for medical diagnostic purposes, when large numbers of eggs exist in the feces sample. The method also recommends examining only a representative amount of the sample collected and averaging over the concentration seen on the main part of the McMaster slide, rather than systematically looking at the entire assay collected. When low numbers of eggs are in the samples, false negative results could be reported. The method uses ether as a reagent which is dangerous to technicians, but does allow for substitution of ethyl acetate which does not affect efficiency and is safer (Ayres et al., 1996).

The 1999 U.S. EPA published method is also proven to detect helminthes, but is more time consuming than the WHO method, especially when a large number of samples are processed. The EPA method also uses ether as a reagent. This method also uses a counting cell, and instructions in the protocol are confusing and incomplete for obtaining representative ‘random’ samples, thus biased counts may occur. Method recoverability of seeded samples was very low for Bean and Brabants (2001), who reported rates of 12% and 4.5% for wastewater and biosolids, respectively (Bowman et al., 2003). There is also a lack of QA/QC data because helminthes are not endemic in the U.S. and routine monitoring is not required (Nelson et al, 2003).

The Tulane method for helminth recovery in wastewater was published in 2001 (ref) The Tulane method is the only method in the literature that has documented recoveries greater than 50% and available QA/QC data for numerous environmental matrices, including alkaline-treated, soil-blended, and lagoon treated biosolids. Spiked sample recoverability of this method is 50% or higher for all of the environmental samples tested. Accuracy and precision for acidic biosolids (pH<11) was 75-80% and 10-15% , and 58-75% and 11-37% for alkaline biosolids. This method uses sieving and flotation steps, eliminating the use of toxic reagents and promoting safer conditions for the laboratory technician. It is also inexpensive, simple, fast, and each of the steps within the procedure have been tested for effect on final recoveries and viability. A larger wastewater sample is taken in this method (19 liters), compared with other methods (1 liter), which is important for samples with low egg concentrations. The final assay collected is examined systematically under the microscope using traditional microscope slides rather than using counting cells. While this method does not have high recovery of very small eggs (Trichuris sp., Taenia sp.) without modification to density/sieving specifications, it will detect the WHO recommended helminth indicator, Ascaris lumbricoides, and other helminth eggs of similar size (Bowman et al., 2003, Nelson et al., 2001). Because of its high recoverability rates and ability to process samples rapidly, this method is the most appropriate for routine monitoring in environmental laboratories of developing countries.

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6.1.3. Ascaris egg recovery with the Tulane Method To test the accuracy of the Tulane Method, biosolid samples (free of helminth eggs) in 19-L water were spiked with 50 or 100 µl of well-mixed positive control Ascaris suum eggs (from pig feces, Excelsior-Sentinel, Inc. (Newfield, NY)). Before spiking the samples, stock egg concentrations were calculated by examining 10 50-µl aliquots of the stock solution. The average concentration and the standard deviation of the stock solution are shown in Table 6.1 below, with the recovery results for the procedure. The recovered concentrations are less than 100% of the spike concentrations, but comparable to recoveries from Bowman et al. (2003). Table 6.1. Tulane Method Ascaris egg recoverability results

Egg Aliquots for Sample Spiking Stock Solution Egg Concentration 50 µl 100 µl Average from 10 counts 230 460 Standard Deviation 15 30 Samples Eggs Recovered % Recoverability 1A—50 µl Spike 117 50.9% 2A—100 µl Spike 360 78.3% 3A—100 µl Spike 273 59.3% 4A—100 µl Spike 305 66.3%

6.1.4. Method Transfer to Mexicali Laboratories To transfer the method to environmental monitoring laboratories in Mexicali, a one-week seminar training session took place in the UABC laboratory with attendees from the university, and the local government agencies (CESPM, CEFPP), including the laboratory technician from the Zaragoza wastewater treatment plant (location of agricultural effluent diversions). Figure 6.1 shows pictures from the training.

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Figure 6.1 Helminth detection method training session at UABC laboratory. 6.2. ILLUSTRATED STEP-BY-STEP HELMINTH DETECTION PROTOCOL IN ENGLISH AND SPANISH The English and Spanish versions of the illustrated step-by-step Tulane method for wastewater and biosolid samples are now given.

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METHOD TO DETERMININE THE CONCENTRATON OF

HELMINTH EGGS IN RAW AND TREATED WASTEWATER—

MODIFIED TULANE METHOD

SCERP PROJECT W-03-13 SEPTEMBER 2004

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METHOD TO DETERMININE THE CONCENTRATON OF HELMINTH EGGS IN RAW AND TREATED WASTEWATER—MODIFIED TULANE METHOD SCOPE Intestinal parasites contaminate the environment when infected human and animal feces are discharged to surface waters or used for agricultural purposes1,2. This document describes the method for recovery, identification, and enumeratation of viable Ascaris spp. eggs in raw and treated wastewater samples. This method has been modified for wastewater from the ‘Tulane’ method used in wastewater biosolid matrices. Endnotes in this protocol provide information for enumeration in sludges and biosolids2. The method is designed to detect Ascaris spp. Because of their similar size and density Toxacara spp. will also be detected by this method. Modifications to the method for detection of Trichuris spp. and Taenia spp. would be necessary because of their greater densities, and smaller sizes. It is possible to recover Trichuris trichiura, but they may pass through the 39 µm sieve on their long axis, which is 20-30 µm. Human hookworm eggs may also be recovered using this method, but identification is difficult due the similarity in morphology and physiology to non-pathogenic helminth eggs. TERMINOLOGY Helminthes—helminthes are a group of organisms that include human and animal intestinal parasites, and non-pathogenic helminthes. Eggs of Ascaris lumbricoides are common indicators of helminth presence. Ascaris spp. eggs have a number of outer layers that aid in identification (Figure 19). The outer-most tanned, bumpy layer is called the ‘mammillated’ layer. Ascaris spp. eggs that do not have this layer are termed ‘decorticated’. If the Ascaris spp. egg is potentially infective, a third-stage larva will be visible within the eggshell after it has developed during incubation (Figure 18) 4.

SUMMARY OF METHOD This method uses a combination of flotation and centrifugation steps to separate the Ascaris spp. eggs from an environmental sample. The eggs are settled from the solution using centrifugation, and washed with a detergent solution to facilitate the separation of them from other solids in the wastewater. The eggs are then floated from the solution and other solids using density-gradient separation, and are captured by sieving the supernatant solutions. For determination of potential infectivity, the isolated eggs are incubated for two to three weeks to embryonate the viable eggs. The isolated eggs are then exposed to a bleach treatment, which decolorizes the outer layer and facilitates the final identification using direct microscopy. REAGENTS, MATERIALS, EQUIPMENT Reagents

1. Anionic Detergent solution: 0.1% (vol./vol.) aqueous solution of Tween 80. Dilute 1 ml Tween 80 to 1000 ml distilled water.

2. Flotation solution. Dissolve 245 g MgSO4-7H2O (USP or ACS, Epson Salt) in 1000 ml of distilled water, for a specific gravity of 1.20. (ZnSO4-7H2O can also be used. Dissolve 330 g in 1000 ml of distilled water, for a specific gravity of 1.2). Measure specific gravity using a hydrometer.

3. Formaldehyde (35%-40% (vol./vol)). 4. Household bleach. 5. Distilled water.

Materials

1. 5-gallon (19 L) straight-sided plastic buckets with lid (See Figure 1 for examples of suitable containers).

2. Centrifuge tubes, conical, 50 ml, polypropylene.

3. Centrifuge tubes, conical, 15 ml, polypropylene.

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4. Microscope cover slips, glass, 24 x 40 mm or 24 x 50 mm.

5. Microscope slides, glass. 6. Sieve, 100 mesh (150 µm). 7.62 cm

diameter (3 in), stainless steel frame. 7. Sieve, 400 mesh (39 µm), 7.62 cm

diameter (3 in), stainless steel frame. 8. Culture dishes, 60 mm glass or plastic

petri dishes. 9. Wash bottles, plastic, 500 ml (Figure

15). 10. Spray bottles, 500 ml (Figure 3). 11. Beakers, 500 ml, glass. 12. Applicator sticks, wooden, 15 cm long.

(Figure 6) 13. Camel hair brush, small. (Figure 11). 14. 50 µl pipette and pipette tips. 15. Parafilm 16. Hydrometer, 1.0-1.3 sp. gr. 17. Fresh Ascaris spp. eggs for positive

control, either dissected from Ascaris suum gravid adult female worms or purified from Ascaris infected pig fecal material.

Equipment

1. Centrifuge, with rotor for 50 ml and 15 ml tubes, capable of 2000 g.

2. Vortex mixer. 3. Compound Microscope, capable of

100X magnification and also between 400X and 1000X magnification.

. Figure 1. Suitable containers for collection and sedimentation of samples. PRECAUTIONS Infective Ascaris eggs can cause disease if ingested. During sample processing latex gloves should be worn. If the positive Ascaris suum

control is harvested from adult worms, a surgical mask and gloves should be worn, or the harvesting should be performed under a biological or chemical safety hood. This is to avoid the development of an allergy to Ascaris pseudocoelomic fluid4. PROCEDURE 1. For raw and treated sewage, take a 5-gallon (19 L) grab sample in a 5-gallon bucket and seal. Transport to the laboratory and settle overnight (or minimum of 3 hours) at ambient temperature. Sample may need to be gently stirred occasionally to help floating solids settle. Laboratory analysis should be performed within 48 hours of sampling. 2. After settling, pour off the supernatant or decant using vacuum without agitating the sediment, and discard (Figure 2). Transfer the remaining sediment to a smaller glass vessel (500 ml) to facilitate pouring into 50 ml tubes, and thoroughly wash all solids from the bucket, using a spray bottle.

Figure 2. Removing supernatant using a vacuum pump and side arm flask. 3. Carefully pour the solution into 50 ml centrifuge tubes, evenly distributing solids (~10 g uncentrifuged solids per tube), and using as many tubes as necessary. Thoroughly wash all solids from beaker into the tubes using a spray bottle (Figure 3). Bring the volume of each tube to 50 ml with distilled water.

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Figure 3. Washing the sediment from sides into tubes with a spray bottle. 4. Centrifuge the tubes for 5 minutes at 2000 g, or maximum speed if the centrifuge maximum speed is less than 2000 g. (Figure 4). Let the centrifuge spin to a complete stop before removing the samples—DO NOT USE THE BRAKE FOR ANY OF THE CENTRIFUGATION STEPS! .

Figure 4. Centrifugation of tubes to settle solids. 5. Pour off the supernatant and discard (Figure 5). The packed sediment should not exceed 5 ml; if so, add water, mix and distribute evenly among additional tubes. Repeat centrifugation for these tubes and decant supernatant.

Figure 5. The supernatant is discarded leaving only the sediment.

6. Wash the sediment by placing 15 ml of detergent solution into the tube and vortexing for 20-30 seconds (the sample can also be mixed by hand). When vortexing, mix with 2 applicator sticks, making sure to dislodge all solids from the bottom of the tube (Figure 6). Set the vortex mixer on a low setting to ensure that solids are not splashed from tube, and direct tube away from body. Once sediment has been resuspended, rinse applicator sticks into tube, and fill the tube to a volume of 50 ml with the detergent solution.

Figure 6. Mixing the sample with a vortex mixer and applicator sticks. 7. Centrifuge the tubes for 5 minutes at 2000 g. 8. Pour off the supernatant and discard. Rewash the sediment by placing 15 ml of distilled water into the tube and vortex for 20-30 seconds with applicator sticks. Bring the tube to a volume of 50 ml with distilled water. 9. Centrifuge the tubes for 5 minutes at 2000 g. (Perform steps 10-14 with 4 tubes at a time, so the eggs are in contact with the flotation solution for the shortest time possible) 10. Add approximately 20 ml of MgSO4 (or ZnSO4) flotation solution, and vortex tubes for 30 seconds with two applicator sticks, mixing thoroughly while vortexing. Rinse applicator sticks and fill tubes to the 50 ml mark with the flotation solution (Figure 7).

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Figure 7. Addition of flotation solution. 11. Centrifuge the tubes for 5-10 minutes at 2000 g. 12. While rotating the tube, pour most of the supernatant (~35 ml) onto two stacked sieves (Figure 8), with the No. 100 sieve stacked on top of the No. 400 sieve. The stacked sieves should be placed over a beaker or small container (Figure 9). Figure 8. While rotating, flotation fluid is poured through stack sieves.

Figure 9. Stacked sieves and beaker set-up

13. Wash the excess fine particles and fluid through both sieves with the wash bottle (Figure 10). A camel hair brush can be used to gently stir the sediment on the surface of the sieves if there is difficulty in passing the supernatant (Figure 11). Up to 4 tubes from the same sample can be poured on the sieves. After each sample, the sieves should be washed with hot

soapy water and a scrub brush to prevent cross contamination.

Figure 10. Rinsing sediment from sieves with wash bottle.

Figure 11. Stirring sediment on surface of sieves using a camel hair brush. 14. The eggs are now captured on the No. 400 sieve. Carefully hold the sieve at a 60o degree angle towards you, and rinse the collected egg solution into the incubation container (petri dish), by directing a stream of water from a wash bottle onto the upper surface of the sieve and thoroughly wash in a downward stream into the petri dish (Figure 12). This should be approximately 10 ml of solution. Make sure that there is enough fluid to cover the bottom of the dish (to a depth of 3-4 mm). If immediate sample processing is required (no viability determination), rinse the sediment into 50 ml disposable cup, as described above, and skip steps 15-16, and 19.

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Figure 12. Transferring eggs from No. 400 sieve to incubation container. 15. Add 100 µl of formaldehyde (per 10 ml of sample) to prevent fungal growth on the container (Figure 13). Place a sheet of parafilm over the petri dish to seal and cover with petri dish lid. Figure 13. Adding formaldehyde to culture dishes for prevention of mold.

16. Incubate for 3 weeks at 22-28oC in the dark or subdued light (Figure 14) (Higher temperatures are better because the larva will develop faster). Reaerate the samples every few days by opening the parafilm and gently agitating for 1 minute.

Figure 14. Incubate samples at 22-28oC in the dark or subdued lig

17. After the incubation period, transfer the cultured eggs to one or more 15-ml centrifuge tubes, by placing a small glass funnel in the tube and rinsing all eggs from the petri dish into the funnel using a wash bottle (Figure 15).

Figure 15. Pouring the egg solution from the petri dish into 15 ml tube using a glass funnel.

18. Centrifuge for 5 minutes at 2000 g. Carefully pipette off supernatant, leaving approximately 0.5 ml of solution at the bottom of the tube. 19. Add 50 µl of household bleach (for a 10% (vol./vol.) bleach solution) and let samples soak for 20 minutes. This decolorizes the outer shell of the Ascaris and Trichuris eggs and allows the inner contents of the egg to be examined more easily during the microscopy analysis. 20. With pipette, mix gently and transfer an appropriate amount (~50 µl) of the egg solution to a microscope slide, and cover with a coverslip (the fluid should not extend beyond the edges of the coverslip). 21. Systematically examine under the microscope at 100X magnification for parasitic helminth eggs (Ascaris spp., Toxacara spp., etc.). If a parasitic egg is found, identify the species using Figures 16-24 under the 400X microscope objective. 22. Count the number of eggs for each type of parasitic helminth egg (Ascaris spp., Toxocara sp., Trichuris spp., etc.) and record whether each is viable (i.e. contains a developed larva) or is non-viable (no larva has developed within the egg). 23. Report the number of viable eggs of each parasitic helminth present per 19 L sample size.

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If no eggs are found, report: >1 viable helminth eggs/19 liters. Identification of Helminth Eggs: In wastewater, parasitic eggs may originate from human or animal origins (i.e. dogs, pigs, rats, birds). By morphological characteristics alone, many of these parasites are almost indistinguishable from human to animal origins (i.e. Ascaris lumbricoides (human origin) and Ascaris suum (pig origins) are morphologically identical, and Toxocara canis (dog origin) are very similar (slightly larger)—refer to Figure 18 and 23). Numerous non-pathogenic helminth eggs and adults also exist in wastewater samples, so identify parasites very carefully to avoid false positives, and do not try to identify adult stages. Look for the distinguishing characteristics in the parasitic organisms, like the 2-3 bumpy, outer layers on the oval Ascaris eggs (Figures 16-20)1. Trichuris trichiura, the human whipworm can be identified through its characteristic barrel shape and two unstained polar plugs (Figure 22). Human hookworm eggs (Ancylostoma duodenale and Necator americanus) are characterized by the oval/elliptical shape and the thin, unstained shell surrounding the larva, but are usually indistinguishable from non-pathogenic eggs (Figure 21). If possible, measure the size of helminth eggs using an eyepiece micrometer, and compare to average sizes listed in the figures. Refer to Ayres and Mara (1996) for instructions on microscope calibration of the eyepiece, available online at: http://www.who.int/water_sanitation_ health/ wastewater/en/labmanual.pdf.

Ascaris lumbricoides eggs, note the distinctive outer layers on the egg

(65 x 50 µm)

Figure 16. Non-viable decorticated Ascaris egg—no

developing larva in the shell6.

Figure 17. Viable Ascaris egg, decorticated,

unembryonated4.

Figure 18. Viable Ascaris egg, decorticated and

embryonated4.

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Figure 19. 2 mammilated, and 1 decorticated Ascaris

eggs6.

Figure 20. Typical microscope view of Ascaris and Trichuris spp. eggs under low magnification (100X) in an environmental sample7.

Other common human intestinal parasites.

Figure 21. Hookworm egg (57-76 x 35-47 µm)6

Figure 22. Trichuria trichuris egg (50-54 x 20-30

µm)6

Figure 23. Toxocara canis (75 x 85-90 µm)6.

Figure 24. Helminth eggs—size comparison picture8. PRECISION AND ACCURACY Accuracy and precision analyses should be performed for every 10th sample, according to the Split/Spike method. To perform the split/spike method, the sample should be divided into 4 sub-samples—three of the sub-samples should be analyzed as usual for precision measurements, and the fourth sub-sample should be spiked with a known amount of Ascaris eggs. Mathematical calculation of the ‘precision’ is the average of the standard deviations for the three sample averages of eggs. ‘Accuracy’ is calculated by the mean percentage of spiked eggs recovered from the fourth sub-sample, according to the following formula2: ( )( )

( )spikeineggssamplesprecisionineggsmeanspikefromeredreeggs

⋅⋅⋅⋅⋅⋅⋅⋅⋅⋅⋅

##cov#

CALIBRATION OF THE CENTRIFUGE To determine the revolutions per minute (rpm) for the relative centrifugal force, the following formula is used:

rRCFkrpm ×

= ,

where,

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RCF = relative centrifugal force (g) r = radius of the centrifuge from the spindle to the center of the bucket (cm) k = 89456 To convert the relative centrifugal force from rpm, use:

( )k

rpmrRCF2

=

ROUTINE MONITORING OF WASTEWATER Routine monitoring of wastewater samples is dependent on the objectives of the project and the type of wastewater treatment. For raw wastewater samples, the concentration of helminth eggs varies significantly over a 24-hour period. It is important to take samples which represent these variations, and it is recommended that a minimum of two composite samples (every hour for a 24-hr period) are taken on days representative for the monthly fluctuations. Wastewater stabilization lagoons consistently remove parasitic helminthes with the proper retention time, even when the plant becomes hydraulically overload or experiences flow variations. The effluent helminth egg concentration should be low and not subject to diurnal or seasonal variations. For each treatment plant that has not been previously sampled, an initial sampling period of several weeks, once or twice a week, in the morning and the evening should be performed. This will provide a good estimate of the long-term quality of the effluent. For restricted and unrestricted irrigation, if proper retention times are met for helminth removal, sampling once a month is recommended. If stabilization lagoons are not the mode of treatment, sampling should be performed weekly1. ACKNOWLEDGEMENTS We would like to thank Dr. Dwight D. Bowman and his laboratory staff at the College of Veterinary Medicine, Cornell University in Ithaca, New York for providing this method, invaluable advice, and laboratory training. This

protocol was collected as part of a Southwest Center for Environmental Research and Policy (SCERP) research grant W-03-13 awarded to Emily Viau and Jordan Peccia at Arizona State University and Socorro Romero at the Universidad de Autonoma Baja California (UABC). The protocol can be viewed in English and Spanish at: http://public. asu.edu/~jpeccia/helminth_protocol.htm. REFERENCES 1. Ayres, R.M., and Mara, D.D. 1996. Analysis of wastewater for use in agriculture: a laboratory manual of parasitological and bacteriological techniques. World Health Organization, Geneva, Switzerland. 2. Bowman, D.D., Little, M.D., and Reimers, R.S. 2003. Precision and accuracy of an assay for detecting Ascaris eggs in various biosolid matrices. Wat.Res. 37:2063-2072. 3. Nelson, K.L., and Darby, J.L. 2001. Inactivation of viable Ascaris eggs by reagents during enumeration. Appl. Env. Micr. 67: 5453-5459. 4. U.S. Environmental Protection Agency. 1999. Control of pathogens and vector attraction in sewage sludge. Environmental Regulations and Technology. Report EPA/625/R-92/013. 5. Parasitology Images, Nematode Parasites, [online]. Available: http://pathcuric1.swmed. edu/MicroBiology/LabRef/Parasites/Nematodes/NematodeTofC.html. 6. Parasite Image Library, Laboratory identification of parasites of public concern. DPDx, [online]. Available: http://www.dpd. cdc.gov/dpdx/HTML/ImageLibrary/Ascariasis_il.asp?body=A-F/Ascariasis/body_Ascariasis_ il_th.htm. Last modified: August, 2004. 7. Helminthes parasitic. Otto Jirovec negatives collection, Dept. Parasitology, UK Prague. [online]. Available: http://parasite.natur.cuni. cz/jirovec/index.php?category=6. 8. Image. Oregon State public health laboratories. [online]. Available: http://www. ohd.hr.state.or.us/phl/imglib/trichuris-ascaris-hookworm.j

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EXAMINATION OF WASTEWATER BIOSOLIDS The procedure for examining wastewater biosolids (sludge) is taken from the ‘Tulane Method’ from Bowman et al. (2003), which also provides recoverability and precision for various biosolid matrices. This protocol differs from the processing of wastewater samples because it requires more initial homogenization and washing steps to detach eggs from the biosolids. ADDITIONAL MATERIALS 1. Sieve, 20 mesh (850 µm). 7.62 cm diameter (3 in), stainless steel frame. 2. Sieve, 50 mesh (300 µm), 7.62 cm (3 in) diameter, stainless steel frame. 3. Blender, glass, capable of holding 1 L 4. Plastic funnel, 150 mm diameter 5. Magnetic stirrer and stir bars 6. Beakers, 1000 ml, tall form, Berzelius. PROCEDURE 1. For liquid and dewatered sludges, take a sample containing 5 g total solids (dry weight basis), place in a blender with 200 ml of distilled water, and blend for 1 minute. Pour sample into a 1000 ml tall form beaker and using a wash bottle, thoroughly rinse blender into beaker. Add 0.1% Tween 80 detergent solution to a concentration of 900 ml, and settle overnight (or a minimum of 3 hours) at ambient temperature. 2. Pour off or vacuum supernatant to just above the layer of solids. Transfer sediment to blender and add water to 300 ml, blend again for one minute at high speed. 3. Transfer to beaker, rinsing blender and add 0.1% Tween 80 detergent solution to reach 900 ml. Settle for two hours at ambient temperature. Pour off or vacuum supernatant to just above the layer of solids. 4. Add 300 ml 0.1% Tween 80 detergent solution, and stir for 5 minutes with a magnetic stirrer.

5. Pour sample through two stacked sieves, with the No. 20 sieve stacked on top of the No. 50 sieve. The stacked sieves should be placed on the plastic funnel sitting on a tall beaker. Rinse the supernatant from the first beaker through the sieves using a camel hair brush to stir the material on sieve, followed by sprayed water. Add 0.1% Tween 80 detergent solution to 900 ml final volume and settle for two hours at ambient temperature. 6. After settling, pour off the supernatant or decant using vacuum without agitating the sediment, and discard. Transfer the remaining sediment to 50 ml tubes, and thoroughly wash all solids from the beaker, using a spray bottle. 7. Centrifuge the tubes for 5-10 minutes at 2000 g. 8. Pour off the supernatant and discard. The packed sediment should not exceed 5 ml; if so, add water, mix and distribute evenly among additional tubes. Repeat centrifugation for these tubes and decant supernatant. 9. Proceed with Steps 9-23 in the wastewater protocol.

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TECNICA PARA DETERMINAR DE LA CONCENTRACION DE HUEVOS DE HELMINTOS EN

AGUAS RESIDUALES CRUDAS Y TRATADAS—

METODO DE TULANE-MODIFICADO

PROYECTO de SCERP W-03-13 SEPTIEMBRE 2004

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TECNICA PARA DETERMINAR DE LA CONCENTRACION DE HUEVOS DE HELMINTOS EN AGUAS RESIDUALES CRUDAS Y TRATADAS—METODO DE TULANE-MODIFICAD0 ALCANCE DE LA TECNICA Parásitos intestinales contaminan el ambiente cuando las heces de humanas y animales contaminadas se descargan a aguas superficiales o se usan en riegos agrícolas1,2. Este documento describe la técnica para recuperación, identificación, y cuantificación de huevos ‘vivos’ de especies de Ascaris en muestras de aguas residuales crudas y tratadas. El método ‘Tulane’ originariamente utilizado para lodos y biosólidos, fue modificado para la identificación de huevos helmintos en aguas residuales. Al final de esta técnica se proporciona información por cuantificación en lodos y biosólidos2. Este método detecta las especies de Ascaris. Las especies de Toxocara también puedan ser detectadas por este método ya que tienen un tamaño y densidad similar al Ascaris. El método debe de ser modificado para la detección de las especies de Trichuris y Taenia por sus altas densidades grandes o sus tamaños pequeños. Es posible recuperar huevos de Trichuris trichiura, aunque puedan llegar a perderse algunos huevos debido a la morfología del huevo porque el eje mayor (20-30 µm) pasa por el tamiz de 39 µm. El gusano de gancho de humanos se recupera también con este método, pero hay dificultad con la identificación debido a la similitud en morfología y fisiología de huevos de helmintos ambientales no patógenos. TERMINOLOGIA Helmintos—Los helmintos son un grupo de organismos que incluyen parásitos intestinales de humanos y animales y helmintos ambientales no patógenos. Huevos de Ascaris lumbricoides son indicadores comunes para identificar la presencia de helmintos. Los huevos de especies de Ascaris tienen un número de estratos exteriores que ayudan a su identificación. La cáscara exterior es bronceada con pozos que se denomina estrato de ‘mammillated’. Huevos de especie de Ascaris que no tienen este estrato se llaman ‘decorticated’. Si los huevos de Ascaris tienen

capacidad de contagio una larva en la tercera etapa sería visible dentro de la cáscara del huevo después de haberse desarrollado en el periodo de incubación4. RESUMEN DEL METODO El método utiliza una combinación de pasos de flotación y centrifugación para separar los huevos de especies de Ascaris de la muestra ambiental. Los huevos se sedimentaron por centrifugación, y se limpian con una solución de detergente para facilitar la separación de éstos del resto del material sólido. Ya que los huevos tienen menor densidad que el resto del material sólido, estos van a flotar en la solución de flotación que seguidamente es tamizado. Para determinar le capacidad de contagio los huevos retenidos por el tamiz, se deben incubar para desarrollar los huevos ‘vivos’. A estos huevos se les realiza un tratamiento de blanqueador para eliminar el color de la cáscara exterior y facilitar también la identificación final con microscopía directa. REACTIVOS, MATERIALES, EQUIPO Reactivos

1. Solución de detergente: 0.1% (vol./vol.) solución acuosa de Tween 80, diluir 1 ml Tween 80 en 1000 ml de agua destilada.

2. Solución de flotación. Disolver 245 gramos de MgSO4-7H2O (Sal de Epson) en 1000 ml de agua destilada, con una gravedad especifica de 1.20. (Alternativamente, se puede utilizar 330 g de ZnSO4-7H2O en 1000 ml de agua destilada, para un gravedad especifica de 1.20). Medir la densidad con un hidrómetro (gravedad especifica del agua es 1.0).

3. Formaldehído (35-40% (vol./vol)). 4. Blanqueador (decolorante de hogar). 5. Agua destilada.

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Materiales 1. Recipientes con lados sin curvaturas de

5-gallones (19 litros) de plástico con tapones. (Figura 1—Ejemplos de recipientes apropiados).

2. Tubos para centrifugadora, cónico, 50 ml, polipropileno.

3. Tubos para centrifugadora, cónico, 15 ml, polipropileno.

4. Cubreobjetos para microscopio, vidrio, 24 x 40 mm o 24 x 50 mm.

5. Portaobjetos para microscopio, vidrio. 6. Tamiz, malla 100 (150 µm), diámetro

7.62 cm (3 in), marco de acero inoxidable.

7. Tamiz, malla 400 (39 µm), diámetro 7.62 cm (3 in), marco de acero inoxidable.

8. Platos de petri, 60 mm, vidrio o plástico. 9. 2 botellas de enjuague, plástico, 500 ml

(Figura 15). 10. Botella rociadota (spray), 500 ml (Fig.

3). 11. Probetas graduadas, 500 ml, vidrio. 12. Barras para agitar de madera, 15 cm

(Figura 6). 13. Pincel de pelo de camello, pequeño

(Figura 11). 14. Pipeta de 50 µl y tapones de pipeta. 15. Embudo, vidrio, diámetro de 50 mm, pie

corto. 16. Parafilm. 17. Hidrómetro (con intervalo de medición

de 1.0 a 1.3 g/cm3). 18. Huevos frescos de especies Ascaris para

control positivo ya sea de un disección de gusano de adulto hembra preñada de Ascaris suum o depurado de heces de cerdo infectados con Ascaris.

Equipo

1. Centrifugadora, con rotores para tubos de 50 ml y 15 ml, vel de rotación de por lo menos 2000 g.

2. Mezclador de vórtice (Agitador de tubos: automático, adaptable, con control de velocidad).

3. Microscopio óptico, aumentos de 100 a 1000X, y platina móvil.

Figura 1. Recipientes apropiados para colección y sedimentación de las muestras. PRECAUCIONES Ingesta de huevos Ascaris puede infectar el analista. Durante el análisis de la muestra, el analista debe utilizar guantes de plástico y una túnica de laboratorio para evitar riesgo de infección. Lavar y desinfectar (con 70% etanol) el área de trabajo, así como el material utilizado por el analista. Si el control positivo de huevos de Ascaris suum es recoger lombrices de adultos, el analista debe utilizar una mascarilla quirúrgica y guantes, o debe preformar la recolección bajo de una capucha de seguridad biológica o química, para evitar alergias debido el liquido de ‘pseudocoelomic’ de Ascaris4. PROCEDIMIENTO 1. Tomar 19 litros en un garrafa de 19 litros (5-galones) de la muestra de aguas residuales, afluente o efluente y cerrar bien. Transportar al laboratorio y sedimentar durante toda la noche (o un mínimo de 3-4 horas) a temperatura ambiente. La muestra debe ser sacudida ligeramente de vez en cuando para ayudar los sólidos flotantes a sedimentar. La muestra debe procesarse dentro de las 48 horas de su muestreo. 2. Después de la sedimentación, extraer a mano o aspirar con vacío el sobrenadante sin agitar el sedimento (Figura 2). Transferir el sedimento a un recipiente de 500 mililitros para facilitar el traspaso a los tubos de 50 ml y debe la garrafa de 19 litros ser enjuagado 2 a 3 veces con agua destilada (botella rociadora).

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Figura 2. Aspirar con vacío el sobrenadante. 3. Con cuidado, verter la solución en approx. 8 tubos de 50 ml (corresponde a approx. 10 gramos de sólidos sin centrifugar por tubo), incluyendo de 2 a 3 enjuagues del recipiente con la botella rociadora (Figura 3). Llenar hasta arriba los tubos de 50 ml con agua destilada.

Figura 3. Enjuagar el recipiente de 500 ml con agua destilada en los tubos de 50 ml. 4. Centrifugar los tubos a 2000 g durante 5 minutos o velocidad máximo si le velocidad de la centrifugadora es menor que 2000 g (Figura 4). Esperar a que la centrifugadora se detenga por completo antes de retirar los tubos. Nota de Importancia: NO INTERRUMPIR LA CENTRIFUGADORA DURANTE SU FUNCIONAMIENTO EN NINGUN CASO!!

Figura 4. Centrifugar para sedimentar los sólidos en los tubos. 5. Remover el sobrenadante (Figura 5). El sedimento no debe ser más de 5 gramos de pelet (sólidos compactos). Así, añadir agua destilada, agitar y redistribuir uniformemente en tubos adicionales. Repetir el paso 4 para estos tubos y decantar el sobrenadante.

Figura 5. Decantar el sobrenadante mientras mantener el pelet en el tubo. 6. Limpiar el sedimento para resuspender el pelet en 15 ml de solución de detergente (0.1% Tween 80) y homogenizar en el mezclador de vórtice durante 20 a 30 segundos en velocidad baja. Cuando se homogeniza—se usa dos barras para agitar para remover los sólidos del fondo del tubo mientras el tubo debe estar alejado del analista (Figura 6). Enjuagar las barras en el tubo y llenar los tubos hasta arriba con la solución de detergente.

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Figura 6. Homogenizar la muestra con un mezclador de vórtice y barras de aplicar. 7. Centrifugar los tubos a 2000 g por 5 minutos. 8. Remover el sobrenadante. Agregar 15 ml de agua destilada para resuspender el pelet agitando con las barras de agitar sobre el mezclador de vórtice durante 20 a 30 segundos en velocidad baja. Diluir los tubos con agua destilada hasta a 50 ml. 9. Centrifugar los tubos a 2000 g por 5 minutos. (Repetir los pasos 10-14 solo con 4 tubos cada vez, porque los huevos deben mantenerse en solución de flotación por el menor tiempo posible.) 10. Añadir approx. 20 ml de solución de flotación MgSO4 (o ZnSO4), y homogenizar con los barras de aplicar y el mezclador de vórtice durante 20 a 30 segundos en velocidad baja. Con la solución de flotación, enjuagar las barras en el tubo y llenar los tubos con solución de flotación hasta arriba (Figura 7).

Figura 7. Añadir de solución de flotación.

11. Centrifugar los tubos a 2000 g por 5 a 10 minutos. 12. Mientras se gira el tubo en un movimiento circular, verter la mayoría del sobrenadante (~35 ml) encima de la estructura de los dos tamices (Figura 8), con el tamiz de malla 100 (150 µl) encima del tamiz de malla 400 (39 µl). Poner un pequeño recipiente debajo de los tamices (Figura 9).

Figura 8. Girar el tubo en círculo mientras se vierte el sobrenadante sobre los tamices.

Figura 9. Estructura de tamices encima del recipiente.

13. Limpiar las partículas excesivas y el líquido a través de ambos tamices con la botella de enjuague (Figura 10). Si se dificultara el pasaje, se puede utilizar un pincel de pelo de camello para remover los sedimentos y sobrenadante sobre la superficie del tamiz (Figura 11). Hasta 4 tubos de la misma muestra se pueden añadir en un mismo juego de tamices. Después del tratamiento de cada muestra, limpiar con agua jabonosa caliente y un cepillo de fregar los tamices para evitar la contaminación de la muestra próxima.

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Figura 10. Limpiar el sedimento a través de ambos tamices con la botella de enjuague.

Figura 11. Darle vueltas en la superficie del tamiz con un cepillo de pelo de camello si hay dificultad en pasar el sobrenadante. 14. Ahora que los huevos están retenidos por el tamiz de malla 400. Llevar con cuidado el tamiz a un ángulo de 60o hacia ti y enjuagar sobre el recipiente de incubación (un plato de petri) (Figura 12). Se deberá obtener una solución de approx. 10 ml. Asegurarse que hay suficiente liquido para cubrir la parte de abajo del plato (al espesor de 3-4 mm). Si es necesario procesar la muestra inmediatamente (sin determinar la viabilidad), enjugar el sedimento sobre la taza disponible de 50 ml, como esta descrito arriba, y saltee los pasos 15-16, y 19.

Figura 12. Transferir del tamiz de Numero 400 al recipiente de incubación (un plato de petri) el completo solución de huevos. 15. Añadir 100 µl de formaldehído por 10 ml de solución para evitar moho en el plato de petri (Figura 13). Sellar el plato de petri con una hoja de parafilm y luego colocar la tapa del plato de petri.

Figura 13. Añadir formaldehído a platos de petri para evitar moho. 16. Incubar durante 3 semanas a 22-28oC en la oscuridad o con mínima luz (Figura 14) (Altas temperaturas llevan a más rápido desarrollo de las larvas). Cada tres días, ventilar las muestras abriendo el parafilm y agitado (con cuidado) por 1 minuto.

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Figura 14. Incubar las muestras a 22-28oC en la oscuridad o luz mínima. 17. Después del periodo de incubación, transferir a uno o más tubos de centrifugación (15 ml) mediante un embudo de vidrio vertiendo completamente el plato de petri y removiendo los restos del plato con una botella de enjuague (Figura 15).

Figura 15. Verter del plato de petri en un tubo de 15 ml con un embudo de vidrio. 18. Centrifugar

los tubos a 2000 g por 5 minutos. Extraer con una pipeta cuidadosamente el liquido sobrenadante hasta tener solo 0.5 ml en el fondo para el posterior análisis. 19. Añadir 50 µl de blanqueador (para un 10% solución de decolorante) y poner en remojo por 20 minutos. Éste elimina el color de la cáscara exterior de los huevos de Ascaris y Trichuris permitiendo fácil examinación del contenido en el interior del huevo durante el análisis en el microscopio. 20. Con la pipeta, mover con delicadeza y transferir un cantidad apropiada (approx. 50 µl) de la solución de huevos a un portaobjeto y cubrir con cubreobjeto (El fluido no debe ir más allá del borde de la hoja de vidrio).

21. Examinar sistemáticamente la muestra de los huevos de helmintos (especies de Ascaris, especies de Toxacara, etc.) con el microscopio bajo el objetivo de 100X. Si encuentra un huevo de helminto, magnificar con el objetivo 400X e identificar con las Figuras 16-24. 22. Contar cada especie de huevos de helmintos (especies de Ascaris, especies de Toxocara, especies de Trichuris, etc.) y registrar si están ‘vivos’ (i.e. larva desarrollada en el interior del huevo) o no ‘vivos’ (larva no desarrollada en el interior de huevo). 23. Expresar los resultados como número de huevos de cada tipo de helmintos ‘vivos’ por 19 litros. Si no se detectan huevos, expresar los resultados como: >1 huevo de helminto ‘vivo’/19 litros. Identificación de los Huevos del Helmintos: En aguas residuales, huevos de parásitos son de origen humano o animal (i.e. perros, cerdos, ratas, o aves). Es importante notar que las características de morfológicas son casi indistinguible entre el origen humano y animal (i.e. Ascaris lumbricoides (origen de humano) y Ascaris suum (origen de cerdo) son idéntico morfológicamente, y Toxocara canis (origen de perro) son muy similar (un poco grande)—ver Figuras 18 y 23). También, en muestras de aguas residuales existen numerosos huevos y gusanos de helmintos que no son patógenos. Por lo que se debe identificar con cuidado los parásitos para evitar resultados positivos falsos y no identificar los gusanos adultos. Observar los características que se pueden distinguir entre los distintos parásitos, como los cáscaras exteriores con pozos en los huevos ovalados de Ascaris (Figuras 16-20)1. Trichuris trichiura (la lombriz de látigo humano) se puede identificar por su forma de barril y dos tapones polares transparentes (Figura 22). Los huevos de lombriz de gancho (Ancylostoma duodenale y Necator americanus) se caracterizan por la forma de ovalo o elíptico y la larva este rodeada de una cáscara delgada transparente, pero los huevos son indistinguible de los huevos de helmintos que no son patógenos (Figura 21). Si

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es posible, medir el tamaño de los huevos de helmintos con un micrómetro de ojo para microscopio, y comparar los tamaños medios con los de las figuras (Remitir a Ayres y Mara, (1996) para instrucciones para la calibración del micrómetro, disponible en-línea a http://www.who int/water_sanitation_health/wastewater/en/labmanual.pdf).

Huevos de Ascaris lumbricoides: observar las cáscaras características en el huevo (65 x 50

µm)

Figura 16. Huevo ‘decorticated’ no ‘vivo’ de Ascaris—no larva desarrollado en la cáscara6.

Figura 17. Huevo viable de Ascaris, ‘decorticated’,

no desarrollado4.

Figura 18. Huevo viable de Ascaris, ‘decorticated’ y

desarrollado4.

Figura 19. Huevos de Ascaris--2 ‘mammilated’, 1

‘decorticated’6.

Figura 20. Vista típica en microscopio de huevos de Ascaris y Trichuris con un aumento de 100X en una

muestra ambiental7.

Otros parásitos comunes de intestinos humanos.

Figura 21. Huevo de lombriz de gancho

(57-76 x 35-47 µm)6

Figura 22. Huevo de Trichuiria trichuris

(50-54 x 20-30 µm)6

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Figura 23. Toxocara canis (75 x 85-90 µm) (especie

de Ascaris de perros)6.

Figura 24. Huevos de Helmintos—figura de

comparación de tamaño8. RECUPERACION Y PRECISION El análisis de recuperación y precisión se debe realizar por cada 10 muestras, según el método de Dividir-Inocular. La muestra se debe dividir en 4 muestras pequeñas—tres muestras se deben analizar normalmente para medidas de precisión, y añadir (inocular) una cantidad conocida de huevos de Ascaris a la cuarta muestra. La ‘precisión’ es el promedio de las desviaciones normales para los promedios de huevos de las tres muestras. ‘Recuperación’ es el porcentaje medio de huevos que se añaden para recuperar la cuarto muestra, con la siguiente fórmula: ( )( )

( )cuartomuestraaanadirhuevosprecisionmuestrasenhuevosmediomuestradehuevos

⋅⋅⋅⋅⋅⋅⋅⋅⋅⋅⋅⋅

##4#

CALIBRACION DE LA CENTRIFUGADORA Para determinar las revoluciones por minuto

(rpm) de la fuerza centrifuga relativa (FCR), la

fórmula es: rFRCkrpm ×

= ,

donde, FCR = fuerza centrifuga relativa (g) r = radio de la centrifuga del distancia al centro al centro del cubo (cm) k = constante cuyo valor es 89456

Para convertir (rpm) en fuerza de centrifuga relativa (RCF), la fórmula es:

( )k

rpmrRCF2

=

RUTINA PARA CONTROLAR AGUAS RESIDUALES La rutina para controlar aguas residuales depende de los objetivos del proyecto y el tipo de tratamiento de aguas residuales. En muestras de aguas crudas (sin tratamiento), la concentración de huevos de helmintos varía considerablemente en un período de 24 horas. Es importante tomar muestras que representen estas variaciones, se recomienda tomar un mínimo de 2 (dos) muestras compuestas (cada hora para 24 horas) mensualmente, en días representativos. Las lagunas de estabilización de aguas residuales que tienen un adecuado tiempo retención, eliminan los huevos de helmintos con regularidad, aunque las lagunas estuvieran sobrecargadas o tuvieran variaciones de flujos. En el efluente de aguas residuales la concentración de huevos de helmintos casi no varía en el tiempo. Por cada planta de tratamiento sin muestreo previo, debe realizarse un período de muestreo de varias semanas, uno o dos veces por semana en la mañana y la noche. Esta proporcionaría una estimación de la calidad del efluente a largo plazo. Para riegos agrícolas limitado y no limitado, si el tiempo de retención es adecuado para la eliminación de huevos de helmintos, se recomienda un muestro mensual. Si no se usa lagunas de estabilización, se recomienda un muestreo cada semana1. RECONOCIMENTOS Agradecemos el Doctor Dwight D. Bowman y su equipo del laboratorio del Colegio de Medicina Veterinaria, Universidad de Cornell, Ithaca, Nuevo York por proporcionar este método, valiosos consejos, y formación de laboratorio. Este protocolo fue completado como parte de una investigación con fondos del Centre Suroeste de Investigación y Política Ambiental (SCERP) Proyecto W-03-13, concedidos a Emily Viau y Jordan Peccia de Universidad de Estado de Arizona (ASU) y

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Maria-Socorro Romero de Universidad de Autónoma Baja California (UABC). El método se puede encontrar en ingles y español a: http://public.asu.edu/~jpeccia/helminth_protocol.htm. BIBLIOGRAFIA 1. Ayres, R.M., y Mara, D.D. 1996. Analysis of wastewater for use in agriculture: a laboratory manual of parasitological and bacteriological techniques (Análisis de aguas residuales para uso en agricultura: un manual de laboratorio de técnicos de parasitiológico y bacteriológico. Organización de Mundo Salud (OMS), Geneva, Switzerland. 2. Bowman, D.D., Little, M.D., y Reimers, R.S. 2003. Precision and accuracy of an assay for detecting Ascaris eggs in various biosolid matrices (Precisión y recuperación de un análisis de detectar los huevos de Ascaris en matrices de biosólidos varios). Wat.Res. 37:2063-2072. 3. Nelson, K.L., and Darby, J.L. 2001. Inactivation of viable Ascaris eggs by reagents during enumeration (Inactivación de los huevos viables de Ascaris por reactivos durante enumeración). Appl. Env. Micr. 67: 5453-5459. 4. EE. UU. Agencia de Ambiental Protección. 1999. Control of pathogens and vector attraction in sewage sludge (Control de la atracción de patogénicos y vectores en lodos residuales). Tecnología y Normas de Ambiental. Informe: EPA/625/R-92/013. 5. Parasitology Images, Nematode Parasites, (Imágenes parasitiológico) [en-línea]. Disponible: http://pathcuric1.swmed. edu/MicroBiology/LabRef/Parasites/Nematodes/NematodeTofC.html. 6. Parasite Image Library, Laboratory identification of parasites of public concern (Biblioteca de Imágenes Parásitos, identificación laboratorio de parásitos de preocupación publico). DPDx, [en-línea]. Disponible: http://www.dpd. cdc.gov/dpdx/HTML/ImageLibrary/Ascariasis_il.asp?body=A-F/Ascariasis/body_Ascariasis_ il_th.htm. Last modified: August, 2004. 7. Helminthes parasitic. Otto Jirovec negatives collection, Dept. Parasitology, UK Prague. [en-línea]. Disponible: http://parasite.natur.cuni. cz/jirovec/index.php?category=6.

8. Imagen. Oregon State public health laboratories. [en-línea]. Disponible: http://www. ohd.hr.state.or.us/phl/imglib/trichuris-ascaris-hookworm.jpg.

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EXAMINACION DE BIOSOLIDOS DE AGUAS RESIDUALES El procedimiento para la examinación de lodos y biosólidos de agua residuales es del ‘Metodo Tulane’ de Bowman et al. (2003). También, el papel proporcionaría el recuperación y precisión de matrices de biosólidos varios. Requiere más pasos de homogenizar y limpiar inicial a la muestra a separar los biosólidos de los huevos. MATERIALES ADDICIONALES 1. Tamiz, malla 20 (850 µm). diámetro 7.62 cm (3 in), marco de acero inoxidable. 2. Tamiz, malla 50 (300 µm), diámetro 7.62 cm (3 in), marco de acero inoxidable. 3. Mezclador, vidrio, capaz de litro. 4. Embudo, plástico, diámetro 150 mm, Nalgene. 5. Mezcladora de barras magnéticos (con agitación) y barras magnéticos. 6. Recipientes, vidrio, 1000 ml, forma talla, Berzelius. PROCEDIMIENTO 1. Para lodos de líquidos y secos, tomar una muestra de 5 g sólidos totales (peso seco), poner en un mezclador con 200 ml de agua destilada, y homogeneizar por 1 minuto a alto velocidad. Verter en un recipiente de 1000 ml y enjuagar 2 a 3 veces con un botella rociadora. Añadir la solución de detergente (0.1% Tween 80) a hasta un volumen de 900 ml. Sedimentar durante toda la noche (o un mínimo de 3-4 horas) a temperatura ambiente. 2. Extraer a mano o aspirar con vacío el sobrenadante sin agitar el sedimento. Transferir el sedimento al mezclador, añadir 300 ml de agua destilada, y homogenizar por 1 minuto a alto velocidad (se enjuaga el recipiente 2 o 3 veces).

3. Transferir al recipiente, enjuagar el mezclador, y añadir la solución de detergente hasta 900 ml. Sedimentar 2 horas a la temperatura ambiente. Extraer a mano o aspirar con vacío el sobrenadante sin agitar el sedimento. 4. Añadir 300 ml de solución de detergente (0.1% Tween 80), y agitar durante 5 minutos con una mezcladora de barras magnéticas. 5. Verter la toda solución sobre la estructura de dos tamices, con el tamiz de malla 20 (850 µm) encima del tamiz de malla 400 (300 µm). Poner un recipiente de 1000 ml y embudo plástico debajo los tamices. Limpiar las partículas excesivas y el líquido a través de ambos tamices con la botella de enjuague. Se puede utilizar un pincel de pelo de camello para remover el sobrenadante sobre la superficie del tamiz. Añadir solución de detergente (0.1% Tween 80) a 900 ml, y sedimentar durante 2 horas a temperatura ambiente. 6. Después de la sedimentación, extraer a mano o aspirar con vacío el sobrenadante sin agitar el sedimento. Con cuidado, verter la solución en approx. 8 tubos de 50 ml (corresponder a approx. 10 gramos de sólidos en el tubo sin centrifugar), incluyendo de 2 o 3 enjuagues del recipiente con una botella rociadora. Llenar los tubos de 50 ml con agua destilada hasta arriba. 7. Centrifugar los tubos durante 5-10 minutos a 2000 g. 8. Remover el sobrenadante. El sedimento no debe ser más de 5 gramos de pelleta. Añadir agua destilada, agitar y diluir uniformemente en tubos adicionales. Repetir pasó 7 para estos tubos y remover el sobrenadante. 9. Aplicar los pasos 9 hasta 23 del método de aguas residuales.

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CHAPTER 7. CONCLUSIONS AND OUTCOMES 7.1. WASTEWATER QUALITY AND AGRICULTURAL REUSE The goal of this research was to assess the current wastewater quality and reuse practice in Mexicali, Mexico to aid in building a reuse program that ensures public health protection and sustainable agricultural practice. The results from this study give the following applicable information and useful tools to the concerned government agencies and agricultural community of Mexicali:

• A template for protecting public health through an evaluative, monitoring, and reuse certification process will help ensure the safety and sustainability of a long-term agricultural reuse program developed for the Mexicali Valley.

• The Zaragoza effluent source is equal to the Colorado River in terms of biological constituents, and consistently meets WHO and Mexican public health standards for unrestricted agricultural irrigation. The potential for secondary contamination in the canal system is minimal because no residences or industrial inputs are in the potential project area.

• In terms of sustainability parameters, the Zaragoza effluent source is similar but of lower quality than the Colorado River. The high salinity of the effluent indicates possible long-term soil salinization and crop problems without a carefully managed irrigation and crop selection program. A large-scale reuse program can cultivate typical Mexicali Valley salt-tolerant crops with assurance of crop yields. Heavy metals accumulation will not present a long-term problem based on the results from this study and Romero et al. (2003).

• The Zaragoza effluent source is higher than the Colorado River in terms of nutrients, thus the fertilizer replacement value will provide a cost benefit with effluent irrigation. Long-term usage may also be beneficial for fertility conditions and improvements to soil properties through nutrient and organic matter inputs (Pescod, 1992).

• Zaragoza Wastewater Treatment Plant constituent removal efficiencies consistently met water quality requirements for agricultural reuse applications, thus no further infrastructure (i.e. disinfection facilities) is needed.

• The Tulane method for detection of Ascaris lumbricoides is most appropriate for environmental monitoring labs.

The reuse of water in agriculture constitutes one of the most valuable tools that

developing countries have to control pollution, recharge aquifers, and to face the challenge of increasing agricultural production with limited water supplies. It is recommended that a 2-year certification of a safe agricultural project area be issued to those areas where Zaragoza effluent will be used for agricultural reuse purposes. No crop restrictions for effluent irrigated fields are required based on the public health conclusions, but the selection of salt-tolerant crops is recommended to ensure crop yield and quality. The irrigation schedule and leaching practices should be based on that used for irrigation with the Colorado River to ensure the long-term sustainability of the agricultural fields, and fertilizer requirements should be adjusted to account for the added nutrients via the wastewater effluent. Mixing the effluent with the Colorado River or irrigating solely with Colorado River water is recommended during the later growing periods

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when there is no fertilizer requirement (i.e. flowering stages) to prevent possible adverse effects in the crop yields. Aquifer water quality monitoring should also be performed to ensure no increases in nutrients and salts.

7.2. NEW RIVER WATER QUALITY—IMPACTS OF AGRICULTURAL REUSE AND IDENTIFIED WASTEWATER FLOWS

Through the wastewater inventory of flows discharged to the New River, critical points in the contamination of the New River were identified to determine opportunities to improve the water quality of this binational river. Conclusions from the water quality trends are:

• The Tula West Drain discharges of industrial and municipal wastewater are the largest source of biological and chemical pollution in the New River, with pesticide and VOC contamination also being evident.

• The major source of salinity to the New River was from its largest flow contributor, agricultural return flows, via the North Collector and Xochimilco Lagoon discharges.

• Trends showed positive water quality impacts to the river from the International Drain, due to Zaragoza effluent discharges and agricultural return flows. The detrimental water quality effects from the Agricola Drain discharges were also dampened with these flow contributions to the International Drain, combined with long travel times to the river.

The water quality model of the New River was used to predict the changes in water

quality due to Zaragoza effluent diversions from the river via the International Drain and the Tula West Drain discharge diversions scheduled upon completion of a new wastewater treatment plant for the Mexicali II service area. From the modeling results, it can be concluded that:

• Effluent reuse in agricultural will have a small negative water quality impact on the New River, while the new WWTP will have a larger positive impact, thus any adverse water quality effects an agricultural reuse program would have on the river system would be compensated for with the removal of major municipal and industrial wastewater flows.

From the literature review of previous studies compared with the findings of this study, the water quality of the New River has significantly improved over the past 20 years, and the substantial clean-up efforts along the main body and connecting drain are evident. Continued efforts to complete the wastewater treatment plant for the Mexicali II Service Area are the most important factor for the overall improvement of the New River water quality. The diversion of effluent for agricultural irrigation is recommended even with the current water quality situation because diversion will have a minimal effect on the river.

7.3. PROJECT OUTCOMES

Dissemination of the project results to aid the development of the Mexicali agricultural effluent reuse program will occur in the following manner:

• A report translated to Spanish will be distributed to the following Mexican government agencies, CNA, CESPM, DDR No. 14, and will include the Ascaris lumbricoides detection protocol previously transferred to the Mexico environmental research laboratories.

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• A presentation in Spanish will be given to the local community, included the relevant government agencies and interested large and small farmers.

• An informational brochure in Spanish will be distributed to the local small and large farmers, detailing the public health results from this study (Appendix C). Currently, the use of Zaragoza effluent is shied away from by the local farmers and agricultural worker due to concerns for public health, so this brochure is intended to persuade farmers of the safety of the effluent to both agricultural workers, and end consumers, including both animals and humans. The brochure distribution is important to promote the use of effluent as an irrigation source water in the upcoming future as urban potable demand increases and potable water sources decrease.

The project results for the New River model will be distributed to the relevant Mexican

agencies, and interested U.S. and California agencies, including parties responsible for the New River within the California Regional Water Quality Control Board and the US EPA. The report in its entirety will be submitted to SCERP and available online, and will be submitted for publication in the Ingeniera Hidráulica de México research journal.