Research Article
1 January 2002

Occurrence of Cryptosporidium and Giardia in Wild Ducks along the Rio Grande River Valley in Southern New Mexico

ABSTRACT

Fecal samples were taken from wild ducks on the lower Rio Grande River around Las Cruces, N. Mex., from September 2000 to January 2001. Giardia cysts and Cryptosporidium oocysts were purified from 69 samples by sucrose enrichment followed by cesium chloride (CsCl) gradient centrifugation and were viewed via fluorescent-antibody (FA) staining. For some samples, recovered cysts and oocysts were further screened via PCR to determine the presence of Giardia lamblia and Crytosporidium parvum. The results of this study indicate that 49% of the ducks were carriers of Cryptosporidium, and the Cryptosporidium oocyst concentrations ranged from 0 to 2,182 oocysts per g of feces (mean ± standard deviation, 47.53 ± 270.3 oocysts per g); also, 28% of the ducks were positive for Giardia, and the Giardia cyst concentrations ranged from 0 to 29,293 cysts per g of feces (mean ± standard deviation, 436 ± 3,525.4 cysts per g). Of the 69 samples, only 14 had (oo)cyst concentrations that were above the PCR detection limit. Samples did test positive for Cryptosporidium sp. However, C. parvum and G. lamblia were not detected in any of the 14 samples tested by PCR. Ducks on their southern migration through southern New Mexico were positive for Cryptosporidium and Giardia as determined by FA staining, but C. parvum and G. lamblia were not detected.
Cryptosporidium and Giardia are enteric parasites that infect a wide range of vertebrate hosts, including birds and mammals. Cryptosporidium parvum and Giardia lamblia are the most common pathogenic species of these parasites in humans. In humans, these organisms can cause persistent diarrhea for 1 to 3 weeks. One of the most common modes of transmission is consumption of feces-contaminated water (17, 25, 34). It is estimated that 80 to 96% of surface waters in the United States are contaminated with Cryptosporidium and Giardia (16, 29). The numbers of waterborne outbreaks of Cryptosporidium and Giardia infections have increased in recent years (23), and there is a potential for large outbreaks, such as that seen in Milwaukee, Wis., in which more than 400,000 people were infected with Cryptosporidium (21). Although Cryptosporidium received a great deal of attention after the Milwaukee outbreak, Giardia continues to be the more common of the two organisms in waterborne disease outbreaks (29).
The contributions of Giardia cysts and Cryptosporidium oocysts from avian species to the concentrations of cysts and oocysts in water samples are largely unknown, as are the extent of transmission of bird-vectored organisms to mammalian hosts and the importance of these parasites in avian species. Two species of Giardia have been isolated from avian hosts; Giardia ardeae has been isolated from great blue herons (9), and Giardia psittacae has been isolated from budgerigars (5, 9). In addition to isolation of Giardia from herons and budgerigars, Giardia has been isolated from straw-necked ibises (12, 24) and parakeets (30). To date, only two valid species of Cryptosporidium (Cryptosporidiumbaileyi and Cryptosporidium meleagridis) have been proven to cause infections in birds (7, 32), and Cryptosporidium has been detected in more than 30 species of birds including geese (Anser anser), tundra swans (Cygnus sp.), black-headed gulls (Larus ridibundus), chickens (Gallus gallus), turkeys (Meleagris gallopavo), mallards (Anas platyrhynchos), and Muscovy ducks (Cairina moschata) (28). However, the majority of avian Cryptosporidium infections have been detected in domestic flocks, and very few studies have examined the occurrence of Cryptosporidium in wild bird populations and in birds during migration (15).
Studies of Giardia and Cryptosporidium have revealed little or no cross-infection of avian and mammalian hosts (11, 19, 27). C. parvum has been shown to cause tracheal infections in inoculated chickens (20), while C. baileyi has been shown to infect humans in a single case (8). Giardia duodenalis (synonyms, G. lamblia and Giardia intestinalis), which commonly infects mammals, has been found to be pathogenic in some birds (35, 36).
Graczyk et al. (15), Eichorst et al. (S. Eichorst, A. Pfeifer, S. Filer, M. Markos, and M. L. Tischler, Abstr. 99th Gen. Meet. Am. Soc. Microbiol., abstr. Q94, 1999), and Harrington et al. (B. J. Harrington, H. Kassa, and M. Bisesi, Abstr. 100th Gen. Meet. Am. Soc. Microbiol., abstr. Q-94, 2000) have demonstrated that Canada geese (Branta canadensis) are able to carry several enteric human pathogens, including G. lamblia, Campylobacter jejuni, and C. parvum. To date, it has not been proven that these organisms replicate in geese, although infectivity experiments have indicated that the pathogens isolated from goose feces can cause infections in mammalian hosts (1315). Harrington et al. (Abstr. 100th Gen. Meet. Am. Soc. Microbiol.) sampled goose feces from 17 sites around the Toledo, Ohio, area and determined that 76.5% of the sites sampled were positive for Cryptosporidium and 17.6% of the sites were positive for Giardia. Graczyk et al. (15) determined that the oocyst concentrations in goose feces ranged from 67 to 686 oocysts per g of feces and the cyst concentrations ranged from 75 to 786 cysts per g.
The duck populations in North America in 1999 and 2000 were the largest since the mid 1950s, with estimated fall migrations of more than 90 million birds (3739). Ducks occupy more diverse aquatic habitats than geese, and their foraging habits may increase the frequency with which they contact waterborne pathogens (6). Ducks also tend to migrate deeper into Mexico and Central America than Canada geese (3, 4). This migration pattern allows ducks to come into contact with poor-quality waters found in Mexico and Central America, which may result in more contact with a wide range of pathogens. Wild ducks have yet to be studied to determine their ability to vector Cryptosporidium or Giardia, although inoculated domestic Peking ducks (A. platyrhynchos) have been shown to vector infectious oocysts (13). Because of the large number of ducks and the diversity of their habitats, it is possible that if ducks do vector Cryptosporidium and Giardia at levels demonstrated in geese, they may have a significant microbiological impact on water quality and in turn on public safety.
The purpose of this study was to determine to what extent Cryptosporidium and Giardia could be isolated from wild ducks during their southern migration through southern New Mexico. Efforts were also made to determine if C. parvum or G. lamblia could be detected in some of the positive samples.

MATERIALS AND METHODS

Study area.

Fecal samples were taken from ducks that were harvested during legal teal season (18 to 24 September 2000) and waterfowl season (18 October 2000 to 21 January 2001). All ducks were harvested on the lower Rio Grande River drainage in southern New Mexico from an area that spans approximately 32 miles from the town of Radium Springs (Dona Ana County, New Mexico) to the town of Vado (Dona Ana County, New Mexico).

Fecal samples.

After harvest, the intestines from below the pancreas to immediately above the anus were removed from the abdominal cavity of each bird. Each sample was placed in an individual Ziploc bag, and the bag was labeled with the species of duck, the date of harvest, and the location. Samples were held for no more than 5 h before further processing. Upon arrival of a sample in the lab, 1 cm was cut from each end of the intestine with a sterile razor blade to eliminate any cross-contamination during processing. The fecal material was then squeezed into a sterile 50-ml centrifuge tube containing 10 ml of a 2.5% (wt/vol) solution of potassium dichromate (K2Cr2O7) to maintain the (oo)cysts during storage. The date of harvest, species of duck, and location were recorded on the tube. Fecal samples were then kept at 4°C for no more than 4 weeks before processing.

Separation of cysts and oocysts from fecal material.

After the samples were removed from storage, they were vortexed thoroughly for 1 min. One-half of each sample (5 to 8 ml) was then sucrose enriched as previously described (26). Briefly, 5 to 8 ml of suspended fecal material was centrifuged (1,100 × g, 10 min). The resulting pellet was washed with 2 to 3 ml of deionized (DI) water and then centrifuged (1,100 × g, 10 min), and the resulting pellet was weighed. The pellet was resuspended in 2 ml of DI water, 2 ml of sucrose (specific gravity, 1.30 to 1.35) was then added, and the solution was gently mixed. The solution was centrifuged at 1,000 × g for 5 min, and the supernatant was removed and placed in a fresh centrifuge tube with 3 volumes of 1× phosphate-buffered saline (PBS) (1.3 M NaCl, 0.03 M KCl, 0.2 M Na2HPO4 anhydrous, 0.01 M KHPO4 [pH 7.4]). The solution was mixed by inverting the tube several times. The (oo)cysts were then pelleted by centrifugation at 2,000 × g for 20 min. The resulting pellet was resuspended in 400 μl of Tris stock solution (50 mM Tris-HCl, 10 mM EDTA [pH 7.2]) and purified further by cesium chloride (CsCl) gradient centrifugation as previously described (33). Three CsCl solutions (solution 1, 1.40 g/ml; solution 2, 1.10 g/ml; solution 3, 1.05 g/ml) were prepared by diluting a stock CsCl solution (1.8 g of CsCl per ml in DI water) with the Tris stock solution. Four hundred microliters of solution 1 was placed in a 2-ml centrifuge tube; solution 1 was then overlaid with 500 μl of solution 2, solution 2 was overlaid with 500 μl of solution 3, and solution 3 was overlaid with the 400-μl (oo)cyst suspension. The gradient was then centrifuged for 60 min at 16,000 × g. Following centrifugation, 920 μl was aspirated from the top of the gradient and placed in a clean centrifuge tube with 1,800 μl of DI water, and the (oo)cysts were pelleted by centrifugation at 5,000 × g for 3 min. The resulting pellet was resuspended in 200 μl of 1× PBS, and (oo)cysts were enumerated by fluorescent-antibody (FA) staining. The remaining fecal sample was retained and used for PCR analysis if necessary.

FA staining.

A cellulose acetate filter (pore size, 0.8 μm; diameter, 13 mm; Sartorius Corp., Filter Division, Hayward, La.) was moistened with 1× PBS and placed into a 13-mm-diameter stainless steel syringe filter holder (Fisher Scientific, Pittsburg, Pa.) attached to a 30-ml syringe clamped to a ring stand. The output of the filter holder was connected to a 250-ml vacuum flask. A sample was added to the syringe, and a vacuum of 2 to 4 in. of Hg was applied to the syringe to pull the sample through the filter. The system was then rinsed with 10 ml of 1× PBS. The filter holder was removed, and 200 μl of antibody (Crypto/Giardia IF test; TechLab, Blacksburg, Va.) diluted 1:10 in a 1% bovine serum albumin solution (1% bovine serum albumin in sterile DI water) was pipetted into the filter holder. Both ends of the holder were covered with aluminum foil, and the sample was incubated for 30 min at room temperature. Following incubation the excess antibody was rinsed from the filter with 15 ml of 1× PBS with a vacuum. The filter was then removed and placed on a glass microscope slide. Five microliters of 1× PBS was placed on the filter, and a glass coverslip was placed on top. The slide was viewed immediately by using epifluorescence. The entire surface of the filter was scanned at a magnification of ×400.

Cell lysis and nucleic acid extraction.

The remaining portions of samples (5 to 8 ml) containing 15 or more (oo)cysts per g of feces were sucrose enriched and centrifuged through a CsCl gradient as described above to collect (oo)cysts for PCR. (Oo)cysts purified by CsCl gradient centrifugation were pelleted by centrifugation (3 min, 5,000 × g). Each pellet was washed once by resuspending it in 1× PBS and centrifuging the preparation for 3 min at 5,000 × g. The pellet was resuspended in 75 μl of Tris-EDTA buffer (10 mM Tris-HCl, 1 mM EDTA [pH 8.0]) with 25 μl of 10% sodium dodecyl sulfate. The solution was then incubated at 37°C for 24 h, and this was followed by phenol-chloroform extraction. The DNA was precipitated by adding 2 volumes of 90% ethanol, 1/50 volume of 5 M NaCl, and 1/250 volume of a 10-μl/ml glycogen solution and incubating the preparation overnight at −20°C. DNA was pelleted by centrifugation (15 min, 21,000 × g). The pellet was washed with 70% ethanol and then resuspended in 30 μl of diethyl pyrocarbonate-treated water (Ambion, Austin, Tex.).

Cryptosporidium PCR assay.

In the amplification assay for Cryptosporidium we used two primer sets. The entire sample (15 μl for primer set 1 and 15 μl for primer set 2) was used for the PCR assay unless the oocyst concentration in the sample was less than 20 oocysts, in which case the entire 30 μl was used with only the species-specific primer set so that an adequate number of targets was present in the reaction mixture. When both oocysts and cysts were present in the same sample, only one Cryptosporidium primer set and one Giardia primer set were used for the sample so that an ample number of targets was present in the reaction mixture.
Primer set 1, which targeted a 256-bp region of the 18S rRNA gene (2), was used to screen for Cryptosporidium oocysts (31). The sequences of the primer set 1 primers were as follows: AW1, TAGAGATTGGAGGTTGTTCCT; and AW2, CTCCACCAACTAAGAACGGCC.
The 100-μl amplification reaction mixture used for primer set 1 contained 10 μl of 10× PCR buffer (GeneAmp; Perkin-Elmer Corp., Foster City, Calif.), 2 mM magnesium chloride, 200 μM (each) dATP, dCTP, dGTP, and dTTP, each primer at a concentration of 0.5 μM, 2 U of DNA polymerase (Amplitaq Gold: Perkin-Elmer), and 15 μl of template. The reactions were performed with a DNA thermal cycler (model 9600 or 4800; Perkin-Elmer). The reaction mixtures were incubated for 7 min at 95°C to activate the enzyme; this was followed by 40 cycles of denaturation at 94°C for 1 min, annealing at 54°C for 1 min, and extension at 72°C for 3 min. A final extension step consisted of 7 min at 72°C and was followed by 5 min of incubation at 4°C to stop the reaction.
Primer set 2, which targeted a 451-bp segment of an undefined region of the Cryptosporidium genome (18), was used to screen for C. parvum (31). The sequences of the primer set 2 primers were as follows: LX1, CCGAGTTTGATCCAAAAAGTTACGAA; and LX2, TAGCTCCTCATATGCCTTATTGAGTA.
The amplification mixture was similar to that described above for primer set 1 except that 3.0 mM MgCl2 was used. The reaction conditions were the same as those described above for primer set 1 except that the annealing temperature was 52°C.

Giardia PCR assay.

In the amplification assay for Giardia we also used two sets of primers (primer sets 3 and 4). For samples positive for Giardia, the entire sample (15 μl for primer set 3 and 15 μl for primer set 4) was used for the PCR assay unless the cyst content in the sample was less than 20 cysts, in which case the entire 30-μl sample was used with only one primer set (primer set 3) so that an adequate number of targets was present in the reaction mixture.
Primer set 3 targeted a 163-bp region of a heat shock protein gene (1). The sequences of the primer set 3 primers were as follows: MA1, AGGGCTCCGGCATAACTTTCC; and MA2, GTATCTGTGACCCGTCCGAG. This primer set detects G. lamblia but has also been observed to amplify Giardia muris (31).
The amplification reaction conditions were similar to those described above for primer set 1 except that 2.5 mM MgCl2 was added to the reaction mixture. The reactions were performed as described above except that the annealing temperature was 55°C.
Primer set 4 targeted a 218-bp segment of the giardin gene (22). The sequences of the primer set 4 primers were as follows: MM1, CATAACGACGCCATCGCGGCTCTCAGGGAA; and MM2, TTTGTGAGCGCTTCTGTCGTGGCGCGCTAA. This primer set has been shown to be specific for G. lamblia (31).
The amplification reaction conditions were similar to those described above for primer set 1, except that 1.5 mM MgCl2 was added to the reaction mixture. The reactions were performed as described above for the other primer sets except that the annealing temperature was 65°C. The optimized conditions for each of the primer sets were described by Rochelle et al. (31). Extracted DNA from purified C. parvum and G. lamblia were used as positive PCR controls for all four primer sets. As a positive purification control, two fecal samples that were negative for both Cryptosporidium and Giardia were spiked with 1,000 purified C. parvum oocysts or G. lamblia cysts. The samples were then purified, the DNA was extracted, and the extract was serial diluted to determine PCR sensitivity.
The PCR products were visualized by electrophoresis on 2% agarose, stained with ethidium bromide, and examined under UV light. A total of 14 samples were screened by PCR.

RESULTS

A total of 69 ducks were sampled during this study. Several species of ducks were sampled, including blue-winged teal (Anas discors), green-winged teal (Anas cercca carolinensis), mallard (A. platyrhynchos), American widgeon (Anas americana), Northern pintail (Anas acuta), hooded merganser (Lophodytes cucullatus), and common merganser (Mergus merganser). Of the 69 ducks sampled, 51 (74%) were mallards, 6 (8.7%) were green-winged teals, 4 (5.7%) were blue-winged teals, 3 (4.3%) were American widgeons, 3 (4.3%) were common mergansers, 1 (1.4%) was a Northern pintail, and 1 (1.4%) was a hooded merganser (Table 1). (Oo)cysts were detected in samples taken during all months of the study (Table 2). The total weights of the fecal samples collected ranged from 0.5 to 3 g per duck. Of the 69 ducks harvested, 49% were positive for Cryptosporidium sp. as determined by FA staining; however, C. parvum was not detected in any of the duck samples when they were tested with a C. parvum-specific PCR primer set. The oocyst concentrations in the samples ranged from 0 to 2,182 oocysts per g of feces, and the mean ± standard deviation was 47.5 ± 270.3 oocysts/g (n = 69). Of the 69 duck samples tested, only 3 contained more than 100 oocysts/g (Fig. 1). The majority of the positive samples contained between 1 and 25 oocysts per g. Twenty-eight percent of the duck samples were positive for Giardia sp., although no G. lamblia was detected in any of the samples by PCR. The cyst concentrations ranged from 0 to 29,293 cysts per g, and the mean ± standard deviation was 436 ± 3,525.4 cysts/g (n = 69). Of the 69 duck samples tested, only 3 contained 100 or more cysts/g (Fig. 1). Most of the positive samples contained between 1 and 25 cysts per g. In 59% of the ducks sampled, either Cryptosporidium or Giardia was present in the feces, while only 14% of the ducks sampled contained both Cryptosporidium and Giardia. Of the 69 samples collected, 14 had (oo)cyst concentrations that allowed screening by PCR. A PCR product was detected only with primers AW1 and AW2 specific for the Cryptosporidium 18S rRNA gene (Table 3). The seven positive samples that tested with this primer set produced strong visual bands, while none of the 10 samples tested positive for C. parvum. All five samples used for Giardia were negative with both primer sets. The levels of sensitivity of the PCR assay for fecal samples were determined to be 10 oocysts for primer sets 1 and 2 and 20 cysts for primer sets 3 and 4.

DISCUSSION

The groups of ducks on the Rio Grande River in the study area can range from small groups consisting of five or six individuals to isolated groups consisting of up to 500 individuals (T. Mitchincin, New Mexico Game and Fish, personal communication). The ducks that migrate through the Rio Grande corridor initiate their migration in southern Colorado or in places as far north as Canada and conclude their migration in places as far south as El Salvador (www.r6.fws.gov/alamosanwr/avian%20page/waterfowl%20banding/waterfow.htm ). Due to the lack of open water in the area around Las Cruces, N.Mex., ducks and other migratory waterfowl become concentrated on the river, producing large flocks of birds during both the fall and spring migrations. Because of the increasing need for communities in the southwest United States to expand their sources of drinking water, it is important to investigate the influence that migratory waterfowl have on surface waters, their potential as vectors for transmission of enteric pathogens to humans and animals, and the impact of these pathogens on migrating ducks. The results of this study indicate that a smaller percentage of wild ducks carry (oo)cysts and that the concentrations are lower than the values reported for Canada geese (15; Eichorst et al., Abstr. 99th Gen. Meet. Am. Soc. Microbiol. 1999; Harrington et al., Abstr. 100th Gen. Meet. Am. Soc. Microbiol. 2000). The mean oocyst concentration (47.5 oocysts/g) that was detected in the ducks was lower than the 67 to 686 oocysts/g reported for Canada geese (15). However, the mean Giardia concentration (436.3 oocysts/g) was in the range reported for Canada geese (15). The mean (oo)cyst concentrations in the duck samples were, however, directly affected by outliers for both the Cryptosporidium- and Giardia-positive samples. If the single duck sample containing 2,182 oocysts per g was removed, the mean oocyst concentration was 16.2 oocysts/g (standard deviation, 72.1 oocysts/g). Removing the single outlier for the Giardia-positive samples (29,293 cysts/g) reduced the mean from 426.3 to 12.1 cysts/g (standard deviation, 55.1 cysts/g).
The high cyst concentration found in a blue-winged teal harvested in September indicated that the duck likely had a Giardia infection at the time that the sample was taken. Giardia infection has not been described previously for wild ducks. Because G. lamblia was not detected by PCR, it is probable that the cysts were cysts of an avian Giardia species. It would be interesting to determine if the Giardia isolated from the ducks is a previously described avian species or if ducks carry a completely new species of Giardia. All of the samples used for Giardia analysis were negative with both primer sets, which indicates that the ducks were not carrying G. lamblia or G. muris but were carrying some other species of Giardia. The sensitivity of the PCR (as determined with oocysts and cysts added to duck feces) indicated that there were enough cysts present that they would be detected if they were amplified with the primer sets (>11 cysts or oocysts/PCR mixture).
Two duck samples contained high oocyst concentrations, which may indicate that replication occurred within the ducks. The PCR data indicates that the oocysts that were isolated from the ducks were oocysts of a species other than C. parvum, probably an avian species of Cryptosporidium or another species of Cryptosporidium that the ducks acquired during foraging. The low numbers of oocysts indicate that most of the positive individuals were not infected. Unlike Canada geese, which have been shown to carry infectious C. parvum (14, 15), wild ducks do not seem to carry these pathogens, at least at levels that can be detected by PCR. Oocyst samples that tested positive for Cryptosporidium did not test positive for C. parvum. However, the numbers of oocysts in most of the positive ducks were low, and only 14 of the 42 positive samples had oocyst concentrations that were above the PCR detection limit. PCR analysis of the samples could not completely rule out the possibility that G. lamblia or C. parvum was present, because the concentration of C. parvum or G. lamblia may have been below the detection limit.
In conclusion, the results of this study indicate that during their southern migration wild ducks do not seem to carry as many Cryptosporidium oocysts as Canada geese carry. However, the Giardia concentrations observed in this study were similar to the Giardia concentrations found in Canada geese (15). The (oo)cysts isolated from the ducks were not C. parvum or G. lamblia (oo)cysts, indicating that the ducks did not appear to be carrying oocysts from the species most commonly linked to human disease. A number of individuals were positive for other species of Cryptosporidium or Giardia, and in a few ducks there was evidence of active replication of non-C. parvum or -G. lamblia isolates, based on the high number of (oo)cysts present.
FIG. 1.
FIG. 1. Distribution of (oo)cyst concentrations in the 69 ducks tested.
TABLE 1.
TABLE 1. Summary of the duck species sampled and the numbers of ducks positive for (oo)cysts
SpeciesCommon nameNo. sampledNo. positive for oocystsaNo. posi-tive for cystsa
Anas discorsBlue-winged teal421
Anas cercca carolinensisGreen-winged teal630
Anas platyrhynchosMallard512313
Anas americanaAmerican widgeon332
Anas acutaNorthern pintail101
Lophodytes cucullatusHooded merganser110
Mergus merganserCommon merganser321
a
Some ducks were positive for both Cryptosporidium and Giardia.
TABLE 2.
TABLE 2. Monthly breakdown of species, number of each species sampled, and number of ducks positive for (oo)cysts
MonthSpeciesNo. sampledNo. posi-tive for (oo)cysts% of ducks positivec
SeptemberaAnas discors3367
 Anas cercca carolinensis31 
OctoberAnas platyrhynchos14747
 Anas discors10 
NovemberAnas platyrhynchos9671
 Anas cercca carolinensis21 
 Anas americana22 
 Lophodytes cucullatus11 
DecemberAnas platyrhynchos17959
 Mergus merganser32 
 Anas cercca carolinensis11 
 Anas acuta11 
JanuarybAnas platyrhynchos11658
 Anas americana11 
a
Only teals could be sampled during the teal season (18 to 24 September).
b
In January samples could be collected on only 21 days.
c
Percentage of all ducks positive for each month.
TABLE 3.
TABLE 3. Comparison of FA staining and PCR results for 14 samples tested by both methods
SpeciesOocysts/gCysts/gPCR results with:
Primer set 1Primer set 2Primer set 3Primer set 4
Anas discors029,293NDaND
Anas platyrhynchos5820+NDND
Anas platyrhynchos250+NDND
Anas platyrhynchos4425NDND
Anas platyrhynchos570+NDND
Anas americana549+NDND
Anas cercca carolinensis2,1820+NDND
Anas cercca carolinensis835NDND
Mergus merganser177NDND
Anas platyrhynchos35146ND
Anas platyrhynchos330+NDND
Anas platyrhynchos280NDNDND
Anas platyrhynchos230NDNDND
Anas platyrhynchos12412+NDND
a
ND, not done.

Acknowledgments

This work was supported in part by the Paso del Norte Health Foundation through the College Project Assistance Initiative (grant 1-4-24503), by the Southwest Consortium of Environmental Research and Policy (grant 1-4-22328), and by the New Mexico Water Resources Research Institute (grant 1-4-23949).

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Published In

cover image Applied and Environmental Microbiology
Applied and Environmental Microbiology
Volume 68Number 1January 2002
Pages: 161 - 165
PubMed: 11772622

History

Received: 11 May 2001
Accepted: 10 October 2001
Published online: 1 January 2002

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Ryan C. Kuhn
Department of Biology, New Mexico State University, Las Cruces, New Mexico 88003
Channah M. Rock
Department of Biology, New Mexico State University, Las Cruces, New Mexico 88003
Kevin H. Oshima [email protected]
Department of Biology, New Mexico State University, Las Cruces, New Mexico 88003

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