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ABSTRACT |
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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A further benefit of such programs is that these regimens result in a significant reduction in prevalence and intensity of intestinal nematode infections, including hookworm (Necator americanus and Ancylostoma duodenale), Ascaris lumbricoides, Trichuris trichuria, and Strongyloides stercoralis. In addition to the well recognized clinical benefits attributable to treatment of these intestinal geohelminth infections, such as the reduction in iron deficiency anemia, and gastrointestinal disturbance, some studies have indicated that such programs result in significant overall improvements in well-being, including school performance and growth among children, outcomes that cannot be accounted for by direct nutritional effects.2,3
With the growing use of broad spectrum anthelmintics, as has occurred with ivermectin in the onchocerciasis control program (OCP), and is now occurring with albendazole in the filariasis elimination programs, an important question emerges as to whether selection pressure will lead to the development of drug resistance. Although resistance to ivermectin has not been reliably documented in the OCP, mathematical modeling indicates that ivermectin distribution will be required for at least an additional 10 years in endemic regions of Africa for effective control.4 While the long life cycle of filarial parasites, and the involvement of an intermediate insect host mitigate against the development of resistance, such a prolonged period of drug use is likely to increase the chances of the development of resistance.4
The potential for the development of resistance among intestinal nematode parasites may be greater than that for filarial parasites: the parasite density is greater in the bowel where the (adult) sexual stages of the parasites reside, and the parasite life cycle is significantly shorter. This is highlighted by experience in veterinary practice, where clinically significant resistance to benzimidazole drugs and ivermectin has emerged.5,6 Three studies have recently been published indicating pyrantel and benzimidazole resistance in human hookworm infection in Africa and Australia.7–9 Although these studies have been criticized for methodologic weaknesses,10 they further highlight the issue of anthelmintic resistance in human gut nematodes.
To lessen the impact of drug resistance, there is a need to develop tools to monitor for its emergence. In this way, selection for resistance may be reduced by, for example, the use of appropriate drug rotation strategies when resistance to one drug group is detected. The fecal egg count reduction test (FECRT), which measures changes in fecal parasite egg counts following chemotherapy, is currently the standard method for determining the therapeutic efficacy of anthelmintic chemotherapy in humans.10 However, studies using this methodology are costly and cumbersome to perform, and are subject to a number of significant confounding factors, including the over-dispersion of parasite burden, leading to sample bias. In veterinary practice, while the FECRT is also the standard method for measurement of resistance, considerable efforts have been made to develop simple in vitro tests as more cost-effective alternatives.11,12 These tests have, in some cases been widely applied in surveys for anthelmintic resistance (e.g., see Palmer and others13). A commercial larval development assay (Drenchrite®; Horizon Technology, Roseville, New South Wales, Australia) is currently marketed in Australia for testing resistance in gastrointestinal nematodes of sheep and goats to benzimidazoles, levamisole, and macrocyclic lactones (MLs), including ivermectin. While such tests for human intestinal nematode infection are conceptually very similar, and a need for their development has been articulated,10 to date little work has been undertaken in this area.
The in vitro techniques for detection and monitoring of anthelmintic resistance in veterinary practice include 1) egg hatch assays, 2) larval paralysis, migration, and motility tests, 3) larval development tests, 4) adult development tests, and 5) metabolic tests.12 While each of these tests has specific strengths and weaknesses, a number of important factors apply in the selection of an assay methodology suitable for human parasites. These include the ability to apply the assay in a field situation with relatively little specialized equipment, a requirement for relatively low numbers of parasites, and, ideally, a method to readily purify parasites from infected stool samples. The aim of this project was to develop an in vitro test for defining drug sensitivity of human hookworm and Strongyloides spp. isolates. The assay was based on assessment of drug effects on the motility of infective-stage larvae. Factors that led us to select an assay based on motility effects included the need for an assay that would be suitable for use in field settings, where the ease of parasite collection from fecal cultures is a major consideration, as well as previous reports of its application to detect resistance to both the benzimidazole and macrocylic lactone drug groups.14,15
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The origin and maintenance of N. americanus and A. ceylanicum in adult laboratory hamsters has been previously described.16,17 Infective larvae of both species were recovered from fecal cultures as described for A. caninum.
For the pilot field study using human hookworm, stool samples were collected from subjects resident in a village in Madang Provence, Papua New Guinea, known to be endemic for N. americanus.18 Fecal samples (approximately five grams taken from fecal collections shown by microscopic examination to be positive for hookworm eggs) were subject to co-proculture using the Harada-Mori technique.19 Briefly, approximately 10 grams of feces was placed on a strip of filter paper and placed in a 50-mL conical tube. Five milliliters of water was added to the bottom of the tube. After 72 hours, the water was poured off and larvae recovered from the fluid by low speed centrifugation (2,000 rpm for 10 minutes). Microscopic examination was undertaken to verify that the species of larvae was hookworm and not the co-endemic nematode Strongyloides fulleborni.
Strongyloides ratti. Since fecal samples from humans with S. stercoralis infection were not available (except for a single case described later in this report), assay development was undertaken using samples from the related rodent species S. ratti. The lifecycle of S. ratti was established in Wistar rats from live infective-stage parasites kindly provided by Dr. Mark Viney (University of Bristol, University, Bristol, United Kingdom). Infective-stage larvae were harvested as previously described.20 Briefly, fecal pellets from infected rats were collected into 9-cm diameter watch glasses and moistened. They were placed into 9-cm diameter square plastic petri plates containing 25 mL of water and maintained at ambient temperature. The water from the petri plates was collected after 5–7 days, and the larvae were harvested and purified as described earlier in this report.
During the course of the study, a single batch of larval isolates of S. stercoralis from an anonymous infected patient was provided by a local diagnostic laboratory. These were collected by agar plate culture.19
Drug sensitivity assay. The assay methodology was adapted from a 96-well microtiter plate assay described by Gill and others15 for measuring the effects of avermectins on the motility of third-stage larvae of the ruminant gut parasite Haemonchus contortus. Stock solutions of ivermectin, thiabendazole, and albendazole (all obtained from Sigma Aldrich, St. Louis, MO) were prepared at 10, 40, or 20 mg/mL, respectively, in dimethylsulfoxide and were serially diluted either two-fold (ivermectin and albendazole) or four-fold (thiabendazole) to produce a series of drug dilutions. Aliquots were added at a dilution of 1% to molten agar in a total volume of 200 µL in individual wells of a 96-well microtiter plate. The final drug concentrations in the assay plates consisted of two-fold serial dilutions starting at 1.6 µg/mL, 0.4 mg/mL, and 0.2 mg/mL for ivermectin, thiabendazole, and albendazole, respectively. Approximately 30 worms (in 30 µL of water) were placed into each well, and the plate was incubated in the dark at 25°C for 48 hours.
The number of separate assay wells used for each series of drug tests varied according to the availability of larvae at different times during the course of the study (see Figure legends and Table footnotes). All assays consisted of at least two wells at each drug concentration (this was increased up to six wells in some cases). Drug sensitivity was generally determined over a series of approximately 10 drug dilutions. However, in some cases, only 4–5 separate drug concentrations were assessed. The number of control wells (no drug added) in each assay varied from 6 up to 21.
The effect of the drugs on worm viability was assessed by counting the numbers of motile larvae after the 48-hour incubation period. Prior to counting, the worms were stimulated to move using hot water as described by Satou and others21 for the assessment of motility in S. ratti and S. venezuelensis following in vitro exposure to drugs. Very little movement was apparent prior to stimulation. However, after the addition of 40 µL of water at 50°C to each well, control worms and those unaffected by drug were observed to move in a rapid sinusoidal motion. In contrast, drug-affected worms showed a twitching motion, or remained motionless. Criteria for distinguishing between motile and non-motile larvae were developed in the first stage of this study, and are described in the Results. Only motile worms were counted in each well. Preliminary experiments showed that the total number of worms in separate wells was almost identical (a repetitive dispensing pipette was used to load larvae into assay plates). Thus, it was considered only necessary to count motile worm numbers to calculate the percent motility relative to a number of control (no drug) wells. The worms were counted under 250x magnification.
Pilot experiments were undertaken to determine the effect on assay results of holding worms for up to five days in water at temperatures of 20, 25, and 29°C prior to undertaking the assay (48-hour assay duration at 25°C), and the effect of varying the temperature that the plates were held at during the 48-hour incubation period (20, 25, and 29°C).
Statistical analyses. Dose-response data was analyzed using non-linear regression (sigmoidal dose-response, GraphPad Prism®; GraphPad Software, Inc., San Diego, CA). Drug sensitivity data were expressed as LC50 values (with 95% confidence intervals). These were defined as the lethal concentrations of drug required to decrease the numbers of motile worms to 50% of that observed in control wells of the microtiter plate.
Ethical approval. Approval for maintenance of the life cycle of S. ratti in laboratory rats was obtained from the Animal Ethics Committee of the Queensland Institute of Medical Research. Necator americanus and A. ceylanicum were maintained in hamsters at the University of Nottingham with the approval of the University of Nottingham Ethics Committee and under a license from the British Home Office. Ethical approval to obtain the infected human feces was obtained from the Human Ethics Committee of the Queensland Institute of Medical Research, and from the Medical Ethics Advisory Committee of the Government of Papua New Guinea.
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RESULTS |
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). A worm was considered to be motile if it moved in a sinusoidal motion when stimulated by water at 50°C. The pattern of the motion differed between the hookworm and Strongyloides species: S. ratti larvae showed rapid motion that could be described as a thrashing motion, whereas A. caninum larvae moved with a smoother, considerably slower motion than S. ratti larvae.
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With all three hookworm species examined (A. caninum, A. ceylanicum, and N. americanus), the distinction between drug-affected non-motile larvae and healthy larvae could be readily scored in the same manner as described earlier for S. ratti larvae exposed to ivermectin. Thiabendazole- and ivermectin-affected hookworm larvae showed very little movement at concentrations slightly higher than the LC50, thus readily enabling the LC50 region to be determined. The degree of movement in thiabendazole- or ivermectin-affected worms was clearly distinguishable from healthy motile worms immediately after addition of hot water to the assay wells. Therefore, it was not necessary to allow a 30-second period to elapse before scoring the motility as was required for S. ratti and thiabendazole.
The assay was unsuitable for examination of sensitivity to albendazole due to the poor solubility of this compound. Precipitated drug was visible in wells containing the drug at concentrations greater than 50 µM. As the drug concentration increased further, the percentage inhibition of motility increased slowly without reaching 100%; at 380 µM, 40% of the A. caninum and 15% of the S. ratti remained motile.
The results of assays performed after a period of 24, 48, or 72 hours incubation are shown in Figure 1. As each assessment required the addition of 40 µL of water, the concentration of drug and larvae was altered, thus precluding repeated assessment of the same assay well at the different timepoints. Therefore, measurements at each timepoint were independently recorded from separate wells. In all cases, as expected, the LC50 decreased as the assay duration increased. The most marked effect of assay duration on LC50 was with the S. ratti thiabendazole combination (Figure 1 A1), where the LC50 after 48 hours of incubation was approximately 8% of the value obtained after 24 hours of incubation. Significantly, for the S. ratti thiabendazole assay, interassay variation in the LC50 was also lower at the 48-hour timepoint compared with the 24-hour timepoint (SE = 46% of the mean at 24 hours versus 33% of the mean at 48 hours). Responses of both S. ratti and A. caninum larvae to ivermectin were more similar at the 48-hour and 72-hour timepoints than for the comparison of 24 hours with 48 hours. Given the finding of reduced interassay variability for both species at an incubation duration of 48 hours, this interval was selected as the standard duration of incubation for future assays.
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. Ancylostoma caninum larvae were significantly more tolerant of storage than S. ratti larvae. Motility of S. ratti larvae decreased significantly when worms were held for three days at either 25°C or 29°C prior to the 48-hour assay (at 25°C) (Figure 2 A1), whereas storage of A. caninum larvae for 5 days at 28°C (prior to a 48-hour assay) had no adverse effects on motility (Figure 2 B1). Despite the markedly decreased motility observed in control S. ratti larvae following storage, the LC50 derived from these assays was similar to that obtained with fresh larvae (up to 1.65-fold less than the LC50 at zero storage time). That is, the LC50 varied little when the number of motile larvae were expressed as a percentage of motile control worms even though the controls were also adversely affected by storage. For A. caninum, storage of worms prior to testing had a similar effect on LC50 (approximately a two-fold variation in ivermectin LC50 after five days storage). Thus, aging of larvae prior to testing did not disproportionately affect the response to anthelmintic.
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). With S. ratti, 2.4- and 1.8-fold decreases in LC50 were observed with thiabendazole and ivermectin, respectively, as the temperature increased from 20°C to 29°C (95% confidence intervals at 20°C and 29°C did not overlap for either drug). The LC50 for ivermectin with A.caninum also showed a similar 1.6-fold decrease; however, the thiabendazole LC50 remained unchanged.
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(insufficient S. stercoralis were available for assays with ivermectin). Typical dose-response curves are shown in Figure 3. There were marked differences in the relative sensitivity of various species to the drugs. Larvae of S. ratti were the most susceptible to thiabendazole and the most tolerant of ivermectin. Necator americanus larvae showed an LC50 to ivermectin approximately two-fold higher than the other hookworm species, and were markedly more tolerant of thiabendazole than all the other species; approximately 50% of N. americanus larvae remained motile at the highest drug concentration that could be examined due to the solubility limitation of the drug (2 mM).
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) with an LC50 value of 0.072, similar to that obtained for the University of Nottingham isolate of this species (Table 3
). Some tolerance was noted when the larvae were assayed with thiabendazole, with up to 28% motility noted in several wells at the highest drug concentration. However, insufficient larvae were recovered from fecal cultures to allow the completion of full dose-response experiments with both drugs.
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DISCUSSION |
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For the assay to demonstrate utility for diagnosis of drug resistance, it will be necessary to establish a correlation between results of the in vitro assay and the sensitivity of a parasite species to a drug in vivo. That is, it will be important to compare in vitro data with therapeutic efficacy as measured by fecal egg count reduction assay, to determine whether differences in the LC50 or LC95 correspond to different rates of parasite clearance. Such a relationship has been established in the veterinary field, where a reduced therapeutic response to ivermectin among certain isolates of H. contortus correlates with a reduced sensitivity to ivermectin among larval stages of the parasite as measured by larval motility.15 This association between in vivo efficacy and the results of a conceptually similar larval development assay has enabled the development and marketing of a commercial test (Drenchrite®; Horizon Technology) to define the resistance status of nematode parasites of sheep and goats towards the benzimidazole, levamisole, and ML drug groups.
The relative tolerance observed in the University of Nottingham N. americanus isolate towards thiabendazole may limit the usefulness of the assay for the detection of resistance to the benzimidazole class of anthelmintics in this species. Only 50% of the larvae were affected by the highest concentration of thiabendazole that could be used in the assay format. Despite this, an apparent a dose-response relationship was evident up to a drug concentration roughly equivalent to the LC50 (Figure 3). Thus, the assay may still be an effective indicator of resistance if the resistance is associated with a significant change in the LC50. While it may be possible to detect benzimidazole resistance in this species by observing a reduction in sensitivity at the highest thiabendazole concentrations, this would represent only the response of the most drug-sensitive individuals among the population. The more tolerant individuals in both susceptible and resistant populations will clearly be unaffected by the drug in the assay. In contrast, for all other drug/parasite combinations examined, a full dose-response relationship over a range up to the complete inhibition of motility was readily demonstrated, and therefore, would enable the detection of changes in LC95. It will be important to determine the dose response towards thiabendazole of N. americanus isolated from humans to assess the potential usefulness of the assay for field measurements of drug susceptibility with this drug/parasite combination.
A variety of in vitro assays have been described for monitoring drug sensitivity of parasites.12 The motility assay described in the present study needs to be examined alongside egg hatch assays and larval development assays to determine the most suitable assay for monitoring the development of resistance in human hookworm and Strongyloides species. Parallel assessments of each assay with susceptible and resistant parasite strains will be required.
The motility assay described here has several advantages for testing of human intestinal nematode drug resistance. These include the requirement of a minimum of fecal manipulations (migration of infective larvae from fecal cultures obviates the need to prepare eggs from feces), and its easily transportable agar matrix format. A significant advantage also is its utility in a field situation where freshly collected human parasites could be studied. Few items of equipment were required (low-speed centrifuge, 250x magnification microscope). The assay temperature and larval storage experiments in this study indicate that variations in these two parameters have little effect on assay results (LC50 varied <2 fold). Ideally, to minimize the effects of assay conditions on dose-response results, parasite larvae isolated in the field would be stored for no longer than one or two days before being assayed at approximately 25–30°C. In addition, the test showed robust performance over a range of temperatures, suggesting that it would work satisfactorily at ambient temperature in tropical environments. A limitation of the initial field testing of this assay was the yield of larvae obtained from the small Harada-Mori cultures. Coproculture of larger samples of feces (complete samples rather than the 5–10 grams used for the present field study) should yield sufficient larvae, as was observed when dog feces were cultured for A. caninum larvae. Once sufficient dose-response data have been collected to define the range of drug sensitivity to be expected in wild-type, susceptible field populations, it may be possible to monitor for resistance using a single discriminating drug concentration rather than depending on a full dose-response assay. This would greatly reduce the numbers of larvae required from each fecal sample.
We observed significant differences in the responses of the five parasite species to the two test drugs, some of which do not correlate with that seen when the drugs are used in vivo. For example, the relative in vitro sensitivity of hookworm and Strongyloides larvae to ivermectin are the reverse of what is seen in vivo. Possible explanations include differences in the mode of action, and physical properties of the drugs resulting in different bioavialability. Although the motility assay has not been evaluated with drug-resistant parasite strains, its ability to discriminate between parasites species and strains showing different responses to drugs suggests its likely utility. For example, N. americanus showed an approximately twofold higher LC50 towards ivermectin than A. ceylanicum, a finding that is in agreement with previous reports22,23 showing that the former species is more resistant to this drug than the latter. However, the 300-fold difference in sensitivity seen in in vivo efficacy trials in the hamster model,22 and the 40–50-fold differences in sensitivity observed in an in vitro motility and ingestion assays using adult worms23 suggests that the larval motility assay may be a less sensitive indicator of drug sensitivity differences between these two species than in vivo or in vitro adult worm assays. However, the basis for differences in drug sensitivity between species may be unrelated to the mechanism of drug resistance shown by different isolates within a species due to the pharmacologic determinants of drug sensitivity, including cuticle penetration rates, detoxification enzyme levels (and substrate specificities), target site sensitivities, and activity of drug efflux pathways. Ultimately, the ability of an assay to detect drug resistance by whatever mechanism that emerges in field isolates of a parasite will determine its utility as a diagnostic test. Thus, assessment of the usefulness of the assay described in the present study for detection of resistance will need to await the examination of resistant strains for each particular species rather than rely on interpretation of current data on species differences.
While current WHO-sponsored efforts to control soil-transmitted helminths are based on the use of albendazole in combination with DEC or ivermectin, we found that albendazole was unsuitable for use in the motility assay. In contrast, thiabendazole performed well in providing clear dose-response data. The inability to directly assess sensitivity to albendazole with this assay may not be an impediment because cross-resistance is the rule within the benzimidazole class of anthelmintics.24,25 Thus, resistance to one drug within the class indicates the existence of resistance to members of the class in general. The suitability of thiabendazole in our assay system identifies it as a suitable drug for use in validating the assay as a drug resistance detection tool for the benzimidazole group of drugs.
While the motility assay has potential as a field-based test for resistance, alternate approaches to resistance detection include polymerase chain reaction (PCR)-based assays. Molecular diagnosis of drug resistance has the potential to detect resistance at much earlier stages compared with in vitro whole-worm assays, thereby allowing appropriate management decisions (drug rotation, etc.) to be made as resistance first appears. Molecular tests for benzimidazole resistance in parasite species of veterinary importance have been described.26,27 Therefore, while the motility assay described in the present study is a potentially useful tool for the detection of resistance, it will also be useful for preliminary validation of PCR tests to relate PCR data directly to drug sensitivity observations.
Received February 3, 2004. Accepted for publication May 11, 2004.
Acknowledgments: We thank Absalom Mai, Melinda Susapu, and Dr. Moses Bockarie (Papua New Guinea Institute for Medical Research, Madang, Papua New Guinea) and Will Kastens and Dr. James Kazura (Case Western Reserve University Center of Global Health and Disease, Cleveland, OH) for their assistance in enabling access to the human clinical samples used to develop the pilot assay, and Dr. Glen Coleman (Veterinary School of the University of Queensland) for undertaking parasitologic analysis on the canine fecal samples.
Financial support: This work was supported in part by grants from GlaxoSmithKline and from the UNICEF/UNDP/World Bank/WHO Special Programme for Research and Training in Tropical Diseases (TDR).
Authors’ addresses: A. C. Kotze and J. O’Grady, Commonwealth Scientific and Industrial Research Organisation Livestock Industries, St. Lucia, Queensland 4067, Australia. S. Clifford and J. M. Behnke, School of Biological Sciences, University of Nottingham, University Park, Nottingham NG7 2RD, United Kingdom. J. S. McCarthy, Queensland Institute of Medical Research, Centre for International and Tropical Health and Nutrition, The University of Queensland, Herston Road, Brisbane, Queensland 4029, Australia, Telephone: 61-7-3845-3796, Fax: 61-7-3362-0104, E-mail: J.McCarthy{at}uq.edu.au.
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| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
), 25°C (
), or 29°C (
) and then placed onto drug assay plates and held for 48 hours at 25°C before assessment of motility in control (A1 and B1) or drug-effected wells of the plate (A2, A3, B2, and B3). For A1 and B1, data shown are the mean ± standrard error, n = 10–21 separate wells. For A2, A3, B2, and B3, data shown are the LC50 calculated using motility data from duplicate or triplicate assay wells at each of 4–5 drug concentrations.
), S. stercoralis (
), and Necator americanus (
) after a 48-hour exposure to thiabendazole or ivermectin. Each data point represents the mean ± standard error of motility scores of three replicate wells at each drug concentration, expressed as a percentage relative to the mean of 9–21 control wells, except for S. stercoralis where each data point represents the mean of duplicate assay wells.