2
Microbe Resistance

OVERVIEW

Since the discovery and subsequent widespread use of antimicrobials, a variety of pathogenic viruses, bacteria, protozoa, and helminths have developed numerous mechanisms that render them resistant to some—and, in certain cases, to nearly all—antimicrobial agents. The focus of this session of the workshop was on exploring some of the latest information emerging about how various important pathogens develop resistance to drugs and how such resistance might be overcome.

The bacterial strains staphylococci, enterococci, and pneumococci pose some of the most serious problems in terms of antimicrobial resistance. Scientists have now acquired detailed information about how these bacteria develop drug resistance. In staphylococci, for example, optimization of resistance depends on the operation of a complex pathway involving a central resistance gene and a number of auxiliary genes. Thus, developing drugs that specifically target any of these genes holds potential for reducing the microbe’s drug resistance. A second novel intervention would target the ecology of these types of bacteria. For example, penicillin-resistant strains of pneumonia bacteria have been found to breed prolifically in the nasopharynx of preschool-age children, particularly those who attend day care centers. Devising interventions to limit antimicrobial exposure might help reduce the genetic propensity of these bacteria to develop drug resistance.

Malaria and schistosomiasis are major health threats in the developing



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2 Microbe Resistance OVERVIEW Since the discovery and subsequent widespread use of antimicrobials, a variety of pathogenic viruses, bacteria, protozoa, and helminths have developed numerous mechanisms that render them resistant to some—and, in certain cases, to nearly all—antimicrobial agents. The focus of this session of the workshop was on exploring some of the latest information emerging about how various important pathogens develop resistance to drugs and how such resistance might be overcome. The bacterial strains staphylococci, enterococci, and pneumococci pose some of the most serious problems in terms of antimicrobial resistance. Scientists have now acquired detailed information about how these bacteria develop drug resistance. In staphylococci, for example, optimization of resistance depends on the operation of a complex pathway involving a central resistance gene and a number of auxiliary genes. Thus, developing drugs that specifically target any of these genes holds potential for reducing the microbe’s drug resistance. A second novel intervention would target the ecology of these types of bacteria. For example, penicillin-resistant strains of pneumonia bacteria have been found to breed prolifically in the nasopharynx of preschool-age children, particularly those who attend day care centers. Devising interventions to limit antimicrobial exposure might help reduce the genetic propensity of these bacteria to develop drug resistance. Malaria and schistosomiasis are major health threats in the developing

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world. Chloroquine was historically the primary drug for treating malaria, but its widespread use has led to increasing microbial resistance. Scientists have now identified a particular type of mutation at a specific location on a single gene as being critical in the development of resistance, and efforts are now under way to develop new drugs that target this resistance mechanism. Praziquantel is the only drug now available to treat schistosomiasis. Since the drug has been in use for more than two decades, concerns are mounting that the parasitic worms that transmit the disease from snails to humans are beginning to become resistant. Among the immediate needs, praziquantel’s effectiveness can be prolonged by more selective use, with treatment targeted only to those people at greatest risk for heavy infection and morbidity, as well as by the use of integrated disease management practices, such as snail control, health education, and improved sanitation. At the same time, new drug development needs to continue in anticipation of the eventual failure of praziquantel efficacy. Influenza is a global threat to health. Vaccines represent the first line of defense against the flu, with a new vaccine being developed and distributed each year in response to the changing genetic composition of the causative virus. Still, vaccines are not a total answer, and several classes of antiviral drugs have been developed to treat infected individuals. Two antivirals— amantadine and rimantadine—have been around since the 1960s. Although effective in some circumstances, both types of drugs suffer from drug-resistance problems. Another family of newer drugs, called neuraminidase inhibitors, shows even more promise, as these formulations appear to pose a reduced risk of triggering resistance. A major problem, however, is that the pharmaceutical companies that produce these newer drugs are not making enough doses to cover medical needs in the event—certain to happen at some point—that a highly modified and virulent form of the influenza virus emerges from the animal world and spreads among the human population worldwide. Adding to concerns about antimicrobial resistance is the possibility that terrorists or a rogue nation might use “bioweapons” to expose large numbers of people to genetically engineered drug-resistant pathogens in order to trigger large-scale disease outbreaks. This scenario was brought into sharp perspective in autumn 2001 by the intentional distribution through the U.S. mail of envelopes containing spores of Bacillus anthracis. One issue considered during this session involved the effects of exposure to both anthrax and ionizing radiation at the same time, conditions that military personnel, in particular, might someday face. Based on a recent study in mice, scientists have been able to identify some fundamental factors that contribute to increased susceptibility to bacterial infections in general, and to B. anthracis in particular, after ionizing radiation, as well as to make some general

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recommendations about effective methods of therapy and prophylaxis following such combined exposures. NEW STRATEGIES AGAINST MULTI-DRUG-RESISTANT BACTERIAL PATHOGENS Alexander Tomasz, Ph.D. The Rockefeller University, New York, NY A major impact of the “chemical warfare” that humanity has been waging against the microbial world on an escalating scale since the discovery of antibiotics is the emergence of a vast variety of resistance mechanisms that have moved into virtually all pathogenic species—viruses, bacteria, and protozoa alike. This emergence has occurred with a swiftness that, on an evolutionary scale, is truly remarkable. The rapid progression from a uniformly antibiotic-sensitive bacterium to a uniformly antibiotic-resistant species is well demonstrated by the case of Staphylococcus aureus, which is a primary agent of hospital-acquired infections. In the early 1940s, when penicillin was introduced into therapy, all strains of staphylococci were highly sensitive to this antibiotic. In less than a decade, S. aureus acquired the penicillinase-based resistance mechanism from an unknown “extra species” source. Penicillin resistance spread across the entire species with the “plasmid epidemic,” and by the late 1950s penicillin was useless against S. aureus. The final stage of this remarkable and sweeping genetic change, propelled by the pressure of antibiotic use, is documented in a recent study conducted in Portugal (Sá-Leão et al., 2001). Screening the S. aureus nasal flora recovered from 1,000 young and healthy volunteers who had never received antibiotics showed that 97 percent of the S. aureus colonizing these individuals produced penicillinase and were resistant to penicillin (Sá-Leão et al., 2001). Clearly, the extra-species drug-resistance gene penicillinase has become a domesticated genetic component of S. aureus without causing any survival deficit to the cells. The penicillin-resistant S. aureus, which was originally associated only with patients in hospitals, has managed to move into the community within 50 years of its appearance on the scene. Equally fast was the response of S. aureus and other staphylococci to the introduction in 1959 of semisynthetic ß-lactam antibiotics, such as methicillin. The first methicillin-resistant S. aureus (MRSA) was detected in the United Kingdom in 1961 (Jevons, 1961). By the 1990s, MRSA had become a globally spread pathogen, making the management of nosocomial S. aureus infections complicated and expensive. Essentially the same phenomena were observed in Streptococcus

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pneumoniae, one of the major community-acquired pathogens of our era. S. pneumoniae are responsible for a series of potentially life-threatening diseases that together cause an estimated 1 million to 3 million deaths worldwide annually. The first penicillin-resistant pneumococci (PRSP) were detected in 1965 (Hansman et al., 1974), followed by increasing numbers of reports on the detection of resistant strains. By the mid-1990s, penicillin-resistant strains had spread globally. In both MRSA and PRSP, antibiotic resistance has unfolded in stages, on a rapid time and geographic scale. Initial detection of MRSA and PRSP was followed by reports on geographic spread. Next came reports on the increase in resistance level and in the frequency of resistant isolates. Eventually, multi-drug-resistant strains carrying resistance traits to different classes of antimicrobial agents also began to appear. In 1993, a small group of international experts, including microbiologists, physicians, and public health personnel, gathered at Rockefeller University for a workshop to survey data on the accelerating spread of multi-drug-resistant pathogens (Tomasz, 1994). By this time, the specter of untreatable bacterial infections had appeared on the horizon as a clear possibility. Strains of common community-acquired and nosocomial pathogens equipped with multi-drug-resistant traits had been identified, with some clinical isolates retaining susceptibility to only a single antimicrobial agent. The workshop participants identified a number of specific genetic events which, if they occurred, could precipitate a genuine public health crisis. Examples of such events include the acquisition of high-level vancomycin resistance among MRSA or pneumococci, and the acquisition of ß-lactamase plasmid by group A streptococci. The alarm sounded at the workshop was recently echoed by the World Health Organization (WHO): “Increasingly drug-resistant infections in rich and developing nations alike are threatening to make once treatable diseases incurable (WHO, 2000). Tables 2-1 and 2-2 illustrate the multi-drug resistance phenomenon: the strikingly successful adaptation of two major human pathogens—S. aureus and S. pneumoniae—to a planetary environment that became saturated with highly toxic substances due to the immense quantities of antimicrobial agents deployed in human and veterinary medicine, in agribusiness, and in virtually the entire biosphere. What can one do in this situation? Clearly the backlash of multi-drug resistance has caught the pharmaceutical chemists and infectious diseases specialists by surprise. In retrospect, it seems that the antibiotic era had two interrelated “cardinal sins.” One sin was the neglect and sometimes complete abandonment of preventive measures in favor of a single-minded antibiotic strategy against bacterial infections. The second was the failure to seriously consider consequences of the fact that the overwhelming major

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TABLE 2-1 Development of Multi-Drug Resistance by S. aureus and S. epidermidis (S: susceptible, R: resistant)   S. aureus ATCC 6538 (1930) MRSA Brazilian epidemic clone (1994) Methicillin-resistant S. epidermidis New York Hospital (1996) Amikacin S R R Amp/Sulbactam - R R Ampicillin S R R Cephalothin S R R Cefotaxime S - - Chloramphenicol S R R Ciprofloxacin S R R Clindamycin S R R Erythromycin S R R Gentamicin S R R Imipenem S R R Oxacillin S R R Rifampin S R R Vancomycin S S S Teicoplanin S S - Tetracycline S R R Trimeth/Sulfa S R - Mupirocine (topical) S R R ity of both the most effective antibiotics and resistance mechanisms are actually products of the microbial world. Antibiotics are produced in tiny quantities and on a microscopic scale by some microbes—presumably for the control of the “quorum” of their habitat—and the producer microbes also invented self-protective resistance mechanisms against their own products. The reintroduction of these highly toxic agents into the biosphere in enormous quantities was a major violation of this quorum sensing. It has amplified local wars among microbes to a global conflict between human and microbe, a chemical warfare in which both offensive and defensive (resistance) armaments came from the microbial world (Tomasz, 2000). The current genomic revolution may offer clues for the production of new antimicrobial agents that would not have been invented by the microbial world during evolution. Development and introduction of such novel agents would be a welcome development indeed. However, it would be naïve to think that the microbial world already “awakened” by the antimicrobial armaments race would simply submit to such new onslaughts. Antibiotic-resistance mechanisms have emerged rapidly in the past, even against completely synthetic agents, such as trimethoprim and fluoro

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TABLE 2-2 Development of Multi-Drug Resistance by S. pneumoniae and Enterococcus faecium (S: susceptible, R: resistant)   S. pneumoniae D39 (1949) S. pneumoniae 6B Dallas, Texas (1992) Enterococcus faecium (VRE) (Tn5482) Memorial Hospital (1996) Amikacin S - R Amp/Sulbactam S R R Ampicillin S R R Cephalothin S - R Cefotaxime S R - Chloramphenicol S R S Ciprofloxacin S R R Clindamycin S R R Erythromycin S R R Gentamicin S - R Imipenem S S R Oxacillin S R R Rifampin S - R Vancomycin S S R Teicoplanin S S R Tetracycline S R S Trimeth/Sulfa S R R Mupirocine (topical) S - - quinolones. Thus, a possible deployment of new antimicrobial agents would not solve the basic dilemma of the antimicrobial armaments race that originates from its erroneous core philosophy: namely, the indiscriminate killing of bacteria by wide-spectrum antimicrobial agents. The fallacies of this philosophy have been pointed out repeatedly (Tomasz, 2000). There are a number of antimicrobial strategies, not yet exploited, that would be more discriminatory and therefore less likely to provoke another wave of drug resistance. The pharmaceutical industry’s traditional approach to drug development has been either to find new wide-spectrum drugs against bacterial targets or to reconfigure old drugs against targets that became inaccessible due to drug resistance. Examples of these two strategies would be the development of new classes of fluoroquinolones and the semisynthetic modification of ß lactams to accommodate the penicillinase-based resistance mechanism. However, there are at least two completely different strategies that offer promise. The first strategy would target the resistance phenotype; the second would target the ecology of resistant bacteria. Recent studies have shown that in both MRSA and PRSP, high-level

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antibiotic resistance requires more than the presence of the central drug-resistance determinant (the mecA gene in the case of MRSA, and the mosaic PBP genes that encode low affinity binding proteins in PRSP). Expression of an optimal high-level antibiotic-resistant phenotype also requires the assistance of a number of additional genetic determinants, the functioning of which is critical for the generation of antibiotic resistance, although the protein products of these genes do not react with the antimicrobial agent. Transposon mutagenesis of a highly methicillin-resistant MRSA strain has identified over 20 such “auxiliary genes” (De Lencastre et al., 1999). Inactivation of these genes had no effect on the transcription of the resistance gene mecA to its gene product (the low affinity penicillin binding protein PBP2A), yet phenotypic resistance of the bacteria was drastically reduced. Recent observations in PRSP identified a similar phenomenon. Inactivation of the small pneumococcal operon murMN, responsible for the production of branched structured components in the bacterial cell wall, caused a complete collapse of the penicillin resistant phenotype in spite of the fact that the primary resistance determinants (the low affinity PBPs) remained unchanged in the mutant bacteria (Filipe and Tomasz, 2000). These observations indicate that reversal of drug resistance is possible by two completely different ways: either by inactivation of the central genetic determinant and its gene product, or by inactivation of the products of auxiliary genes (see Figure 2-1). It follows that auxiliary genes represent novel types of antibacterial targets. Compounds capable of inactivating the products of these genes should represent synergistic agents that together with ß-lactam antibiotics would render resistant bacteria sensitive again to these classical antimicrobial agents. FIGURE 2-1 Two ways to reverse drug resistance.

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A major roadblock to development of such agents, however, is that this approach seems to collide head-on with the central philosophy of the pharmaceutical industry, which is only willing to invest in the development of wide-spectrum antimicrobial agents that can assure a market in the range of $1 billion a year. Clearly, the types of agents described here would be specific for the particular bacterial pathogen and therefore would be outside such marketing interest. While the position of “big pharma” on this issue is based on complex economic realities, I believe that the future points in a different direction: the development of highly specific narrow-spectrum agents, the deployment of which would not challenge the entire microbial world each time they are used in therapy. Such development will be hastened by current progress in devising highly sensitive molecular techniques for rapidly detecting and identifying bacterial pathogens—a capability that may lie in the not too distant future. With rapid and safe diagnostics at hand, the use of wide-spectrum antimicrobial agents should be reserved to special cases only because of their indiscriminate challenge to both harmful and harmless bacteria. A second novel intervention with bacterial pathogens, particularly drug-resistant strains, would target the ecology of these bacteria. Recent observations indicate that the overwhelming majority of diseases caused by resistant strains of S. aureus are linked to a surprisingly few epidemic clones or genetic lineages that have immense geographic spread (Oliveira et al., 2002) and that appear to combine in their genetic backgrounds not only determinants of antibiotic resistance but also genes that assure ecological success (i.e., spread and colonization) of the bacteria (see Figure 2-2). Similar observations also have been made for penicillin-resistant S. pneumoniae (Sá-Leão et al., 2000). Clearly, identification of determinants of epidemicity may provide completely new targets—vaccines or chemical agents—against specific multi-drug-resistant clones that are responsible for most of the hardships of resistant disease. Following up on this ecological reasoning raises questions related to the ecological reservoirs of bacterial pathogens, particularly the drug-resistant clones. It has been clearly shown that in the case of PRSP a major sanctuary and breeding ground of drug-resistant strains is the nasopharynx of pre-school-age children, particularly those who attend day care centers. All children have immature immune systems. When this natural condition is combined with the close contact among children that is characteristic of day care centers, the high frequency of viral respiratory diseases in such centers, and the use (and misuse) of immense quantities of antimicrobial agents, the result is the creation of a bona fide “factory” of resistant pneumococci (Sá-Leão et al., 2000). Similar studies could not identify a comparable reservoir of MRSA among healthy carriers (Sá-Leão et al., 2001).

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FIGURE 2-2 Geographic spread of pandemic MRSA clones. Rather, it seems that for resistant staphylococci the ecological reservoir is the hospital itself. A novel and potentially effective intervention to reduce the spread of resistant forms of these two important pathogens would involve intervention at the level of their ecological reservoirs—namely, lowering the carriage rate of resistant bacteria. The European Community has recently initiated such a major project (EURIS, European Resistance Intervention Study), which is aimed at identifying the most effective intervention strategies by which carriage of resistant pneumococci could be reduced among children attending day care centers in member countries (Sá-Leão et al., 2000). An analogous attempt for MRSA would zero in on the hospital itself by introducing rigorous infection-control measures, such as those that have been successfully tested and advocated by several recent studies (Farr and Jarvis, 2002; Pittet, 2002). MALARIA AND THE PROBLEM OF CHLOROQUINE RESISTANCE Thomas E. Wellems, M.D., Ph.D. Laboratory of Malaria and Vector Research National Institute of Allergy and Infectious Diseases National Institutes of Health, Bethesda, MD The discovery of chloroquine nearly 70 years ago had a considerable impact against the morbidity and mortality of malaria. This impact had

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such effect that chloroquine became recognized as one of the most successful and important drugs ever deployed against an infectious disease. Massive use of the drug, however, eventually produced resistant malaria strains (Peters, 1989). First reports of chloroquine resistance were with Plasmo dium falciparum, the species responsible for the most acute and deadly form of human malaria (Payne, 1987). By the 1970s, resistant P. falciparum strains were established in South America, India, Southeast Asia, and Papua New Guinea. Africa was spared until the late 1970s, when resistance was detected in Kenya and Tanzania, seeding the spread of resistance across the continent within a decade (Peters, 1987). In the absence of a replacement drug with the low cost and reliability of chloroquine, the morbidity and mortality from malaria resurged in Africa (Greenberg et al., 1989; Trape et al., 1998). Molecular Basis of Chloroquine Action and Resistance Chloroquine interrupts hematin detoxification in malaria parasites as they grow within their host red blood cells (Chou et al., 1980) (see Figure 2-3). FIGURE 2-3 The pathway of hemoglobin digestion and hematin polymerization in a P. falciparum-infected red blood cell. Chloroquine accumulates in the acid digestive food vacuole of sensitive parasites and interferes with polymerization. Chloroquine-resistant parasites reduce this accumulation and thereby reduce drug toxicity.

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Hematin, a toxic ferriprotoporphyrin product released from digested host hemoglobin, is normally detoxified in the parasite’s acid food vacuole by polymerization into innocuous pigment crystals (Dorn et al., 1998). Chloroquine interferes with polymerization and poisons the parasite by complexing with hematin and adsorbing to the growing faces of the crystals (Sullivan et al., 1996; Pagola et al., 2000). Chloroquine-resistant P. falciparum survives drug exposure by reducing the accumulation of chloroquine in the digestive food vacuole (Verdier et al., 1985). The mechanism of this reduction, not yet established, may involve changes in digestive vacuole pH or a direct effect on drug flux across the digestive vacuole membrane. Chloroquine resistance results from multiple mutations in PfCRT, a P. falciparum protein located at the parasite’s digestive vacuole membrane (Fidock et al., 2000). PfCRT contains 10 predicted transmembrane segments and has a structure consistent with a transporter or channel (Nomura et al., 2001) (see Figure 2-4). Although the exact patterns of PfCRT mutations differ according to the geographic origin of chloroquine-resistant parasites, all of these patterns include a key substitution for lysine at position FIGURE 2-4 Schematic representation of PfCRT and positions of mutations associated with chloroquine resistance. The critical K76T mutation occurs in the first of the ten predicted transmembrane domains. Filled circles show the positions of all other PfCRT mutations that have been identified in different chloroquine-resistant isolates. These mutations may compensate for the K76T change or help maintain critical functional properties of the PfCRT molecule in resistant parasites.

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together with basic clinical support are fundamental. Quinolones have been recommended for preventing sepsis by selective decontamination of the intestinal tract. Although anti-gram-positive antibiotics could be used to supplement other antimicrobial agents, it is imperative that the anaerobic microflora in the intestine not be suppressed because ionizing irradiation reduces them by several logarithms and they are required to provide colonization resistance. Selected non-specific biological response modifiers (BRMs), which enhance innate immunity by inducing cytokines naturally, or specific BRMs, such as cytokines, could augment specific antimicrobial agents, but this combined approach to therapy following irradiation remains essentially experimental (Peterson et al., 1994). Probiotics, such as specific strains of Lactobacillus, might also offer an advantage by enhancing colonization resistance and restoring the intestinal microflora. Factors that influence therapy for post-irradiation infection include: (i) reduced innate immune responses, particularly the decreased number of phagocytic cells; (ii) the pathogenesis of microorganisms; (iii) coverage by selected bactericidal (not bacteriostatic) antimicrobial agents, which cover facultative gram-positive and gram-negative (but not anaerobic) bacteria; (iv) the half-life of the selected antimicrobial agent (which in turn determines the dose schedule); (v) the route of administration (oral is optimal following irradiation, especially for large numbers of casualties); and (vi) the starting time and duration of antimicrobial therapy. The general principles of pharmacokinetics and pharmacodynamics must be considered as well. Figure 2-9 shows the concept that the concentrations of the selected antimicrobial agents must remain above the threshold minimum bactericidal concentration to achieve successful therapy after irradiation because of the absence of an effective innate response and to maintain as high an area under the curve divided by the MBC (AUBC) as practical. That is, in principle, the higher the AUBC, the greater the cidal effect against the bacteria. Selection of an effective antimicrobial chemotherapeutic regimen also depends upon (i) the cidal mechanisms of action of the selected agents, whether by inhibiting formation of cell wall, interrupting cell membrane function, interfering with DNA function or replication, inhibiting protein synthesis, or antagonizing metabolism, and (ii) microbial drug resistance of bacteria, whether by selection of a pre-existing genetic ability in a population or by mutation, which changes the genetic ability of the microorganism. For antimicrobial therapy for sepsis after irradiation, second-generation quinolones, either ciprofloxacin or levofloxacin, are recommended as the first choice, third- or fourth-generation cephalosporins, either ceftriaxone (third-generation) or cefepime (fourth-generation) as a second choice, or aminoglycosides, either gentamicin or amikacin, as a third choice,

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FIGURE 2-9 Concept of the relationship between pharmacokinetic factors, minimum inhibitory concentration (or minimum bactericidal concentration) of an antimicrobial agent and the area under the curve versus time after administration of the drug. with or without amoxicillin or vancomycin as an adjunct, for a duration of 21 days (Brook and Ledney, 1992). Experimental Antimicrobial Therapy for B. anthracis Sterne-Induced Polymicrobial Sepsis After Irradiation The currently recommended treatment for anthrax for up to 60 days is the quinolone, ciprofloxacin i.v., penicillin G i.v., or the tetracycline, doxycycline i.v. (Dixon et al., 1999; Inglesby et al., 1999). Figure 2-10 depicts our experimental design for evaluating antimicrobial therapeutic agents against B. anthracis Sterne infection in sublethally irradiated (7 Gy) B6D2F1/ J female mice. Antimicrobial agents were given for 7, 14, or 21 days after intratracheal spore challenge (Elliott et al., 2002). To determine the effect of starting time of therapy on survival, irradiated mice, 12 per group, were given 7.8 × 108 CFU B. anthracis Sterne spores i.t. Penicillin G, 62.5 mg i.m., was started 6, 24, or 48 hours after spore challenge and continued for 7 days through day 11. Survival was prolonged when penicillin was started 6 or 24 hours after challenge, but when penicillin G therapy was delayed for 48 hours, survival was essentially the same as the control (Elliott et al., 2002).

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FIGURE 2-10 An experimental design for evaluating antimicrobial therapeutic agents against B. anthracis Sterne infection in sublethally irradiated (7 Gy) B6D2F1/ J female mice. SOURCE: Elliott et al., 2002. To compare survival following therapy with penicillin and the two quinolones, ofloxacin and trovafloxacin, irradiated mice, 19 or 20 per group, were given 4.1 × 108 CFU B. anthracis Sterne spores i.t. The quinolone ofloxacin, 40 mg/kg p.o., and penicillin G, 125 mg i.m., were given either separately or in combination, and the quinolone, trovafloxacin, 20 mg/kg, was given either s.c. or p.o. Administration of the agents was started 24 hours after i.t. spore challenge. No control mice survived. Survival was 20 and 25 percent in mice given ofloxacin or penicillin G separately. When these two agents were combined, survival was increased to 55 percent. Survival in mice that were given trovafloxacin p.o. or s.c. was 95 and 100 percent, respectively (Elliott et al., 2002). To evaluate efficacy of macrolides against B. anthracis infection, irradiated mice, 20 per group, were given 1.8 × 108 CFU B. anthracis Sterne spores i.t. Mice were given doses of macrolides s.c. or p.o., which were based on allometric scaling to 10 times the equivalent doses in humans, starting 24 hours after i.t. spore challenge for 14 days through day 18. Sterile water was given either s.c. or p.o. to control groups of mice. Azithromycin (AZM, 50 mg/kg) was given either s.c. or p.o. Clarithromycin (CLR, 150 mg/kg) and erythromycin (ERY, 500 mg/kg) were given only p.o., and the quinolone, trovafloxacin (TVA, 20 mg/kg) was given p.o. for comparison with previous results and as a sort of “positive” control. Only one of 40 water-treated control mice survived. Few mice that were given the macrolides survived (between 0 and 15 percent). Survival was 80 percent in mice given trovafloxacin for comparison. The macrolides may be ineffective because they tend to accumulate in tissues and, so, concentrations in serum are low (Elliott et al., 2002).

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Antimicrobial Resistance in B. anthracis Sterne in Vitro We have not observed a change in antimicrobial susceptibility in B. anthracis Sterne against penicillin G, ciprofloxacin, levofloxacin, and vancomycin during the course of 21 days of antimicrobial therapy. However, we evaluated the potential for B. anthracis Sterne to develop antimicrobial resistance in vitro by passing growth of the bacteria sequentially in minimally inhibiting concentrations of quinolones and doxycycline. Each drug was diluted two-fold as depicted in Figure 2-11. A 0.1-ml amount of a suspension of bacteria, which contained approximately 2.0 × 107 CFU B. anthracis Sterne, was added to each tube in the series and allowed to incubate at 35°C. The MIC was determined by visual observation of microbial growth at 24 and 48 hours. The highest subinhibitory concentration of each antimicrobial agent in which microbial growth was observed was then used as the inoculum for the next series of dilutions. This macrodilution method (Davies et al., 1999) was performed in duplicate with each antimicrobial agent for 21 serial passages to simulate 21 days of therapy. The graph in Figure 2-12 shows the results from the evaluation with alatrofloxacin, a prodrug of trovafloxacin. The MIC began to increase four times or greater than the initial MIC by the ninth passage. So, it appears that a subpopulation of B. anthracis Sterne possesses the propensity to develop resistance against alatrofloxacin in vitro. Results with ciprofloxacin, FIGURE 2-11 Two-fold macrodilution of an antimicrobial agent in vitro to determine minimum inhibitory concentration (MIC) of the drug against a strain of bacteria.

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FIGURE 2-12 Change in minimum inhibitory concentration of alatrofloxacin against Bacillus anthracis Sterne in brain heart infusion during the course of 21 serial passages of the microorganism performed in duplicate. SOURCE: Elliott, unpublished data; Brook et al., 2001b. gatifloxacin, and ofloxacin were similar but we observed only a minimal increase of the MIC for doxycycline. Cross-susceptibility was also determined with the cultures from the twenty-first passage among the quinolones. All substrain isolates that were grown in the presence of one quinolone were also resistant to the other quinolones (Brook et al., 2001b). Antimicrobial agents alone are not likely to resolve infection by B. anthracis, particularly following irradiation, because pathogenesis and death from B. anthracis is mediated by lethal and edema toxins together with a polymicrobial sepsis. Therefore, once the toxins are formed, antimicrobial agents alone are not adequate to prevent mortality. By extending the general approach to treating sepsis following irradiation, successful management of anthrax following irradiation will include the following elements discussed below. New quinolones are effective against both B. anthracis and endogenous bacteria. Agents with a wide spectrum of activity and high concentration in serum are more effective than agents with limited spectrum (e.g., penicillin and early quinolones) or low serum concentration (e.g., macrolides). Bacil lus anthracis could develop antimicrobial resistance with prolonged therapy. Vaccination early during therapy or injection of anti-serum could be a valuable adjunct to inactivate lethal and edema toxins. The standard, initial vaccination requires three injections two weeks apart. Horse anti-serum was used during the treatment of victims of the release of virulent B.

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anthracis spores in Sverdlovsk, USSR, in 1979, but the patients died, so anti-serum seems to have limited value (Abramova et al., 1993). Summary We demonstrated (i) that low-level, acute radiation combined with endemic and BW agents, B. anthracis in particular, increases mortality synergistically; (ii) improved, effective therapy, that is, recent quinolones, to control mixed, polymicrobial infection with intestinal bacteria, which is induced by B. anthracis spore challenge after irradiation; and (iii) development of antimicrobial resistance in vitro in B. anthracis Sterne. REFERENCES Abramova FA, Grinberg LM, Yampolskaya OV, Walker DH. 1993. Pathology of inhalational anthrax in 42 cases from the Sverdlovsk outbreak of 1979. Proceedings of the National Academy of Sciences90:2291–2294. Babiker HA, Pringle SJ, Abdel-Muhsin A, Mackinnon M, Hunt P, Walliker D. 2001. High-level chloroquine resistance in Sudanese isolates of Plasmodium falciparum is associated with mutations in the chloroquine resistance transporter gene pfcrt and the multi-drug resistance gene pfmdr1.Journal of Infectious Diseases183:1535–1538. Baird JK, Leksana B, Masbar S, Fryauff DJ, Sutanihardja MA, Suradi, Wignall FS, Hoffman SL. 1997. Diagnosis of resistance to chloroquine by Plasmodium vivax: timing of recurrence and whole blood chloroquine levels. American Journal of Tropical Medicine and Hygiene56:621–626. Basco LK and Ringwald P. 2001. Analysis of the key pfcrt point mutation and in vitro and in vivo response to chloroquine in Yaoundé, Cameroon. Journal of Infectious Diseases 183:1828–1831. Bennett JL, Day T, Liang FT, Ismail M, Farghaly A. 1997. The development of resistance to anthelmintics: A perspective with an emphasis on the antischistosomal drug praziquantel. Experimental Parasitology87:260–267. Brindley PJ. 1994. Drug resistance to schistosomicides and other anthelmintics of medical significance. Acta Tropica56:213–231. Brook I and Elliott TB. 1991. Quinolone therapy in the prevention of mortality after irradiation. Radiation Research128:100–103. Brook I and Ledney GD. 1992. Quinolone therapy in the management of infection after irradiation. Critical Reviews in Microbiology18:235–246. Brook I, Elliott TB, Harding RA, Bouhaouala SS, Peacock SJ, Ledney GD, Knudson GB. 2001a. Susceptibility of irradiated mice to Bacillus anthracis Sterne by the intratracheal route of infection. Journal of Medical Microbiology50:702–711. Brook I, Elliott TB, Ledney GD. 1999. Infection after ionizing irradiation. In Zak O, Sande MA, eds. Handbook of Animal Models of Infection: Experimental Models in Antimicro bial Chemotherapy.San Diego: Academic Press. Pp. 151–161. Brook I, Elliott TB, Pryor HI 2nd, Sautter TE, Gnade BT, Thakar JH, Knudson GB. 2001b. In vitro resistance of Bacillus anthracis Sterne to doxycycline, macrolides, and quinolones. International Journal of Antimicrobial Agents18:559–562. Brook I, Walker RI, MacVittie TJ. 1988. Effect of antimicrobial therapy on bowel flora and bacterial infection in irradiated mice. International Journal of Radiation Biology53:709– 716.

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Carlton JM, Fidock DA, Djimdé A, Plowe CV, Wellems TE. 2001. Conservation of a novel vacuolar transporter in Plasmodium species and its central role in chloroquine resistance of P. falciparum. Current Opinion in Microbiology4:415–420. Chan MS. 1997. The global burden of intestinal nematode infections: 50 years on. Parasitol ogy Today13:438–443. Chen N, Russell B, Staley J, Kotecka B, Nasveld P, Cheng Q. 2001. Sequence polymorphisms in pfcrt are strongly associated with chloroquine resistance in Plasmodium falciparum. Journal of Infectious Diseases183:1543–1545. Chou AC, Chevli R, Fitch CD. 1980. Ferriprotoporphyrin IX fulfills the criteria for identification as the chloroquine receptor of malaria parasites. Biochemistry19:1543–1549. Cioli D. 1998. Chemotherapy of schistosomiasis: an update. Parasitology Today14:418–422. Cioli D and Pica Mattoccia L. 1984. Genetic analysis of hycanthone resistance in Schistosoma mansoni. American Journal of Tropical Medicine and Hygiene33:80–88. Cooper RA, Ferdig MT, Su X, Ursos LM, Mu J, Nomura T, Fujioka H, Fidock DA, Roepe PD, Wellems TE. 2002. Alternative mutations at position 76 of the vacuolar transmembrane protein PfCRT are associated with chloroquine resistance and unique stereospecific quinine and quinidine responses in Plasmodium falciparum. Molecular Pharmacol ogy61:35–42. Cowman AF, Karcz S, Galatis D, Culvenor JG. 1991. A P-glycoprotein homologue of Plas modium falciparum is localized on the digestive vacuole. Journal of Cell Biology 113:1033–1042. Cravo P, Culleton R, Hunt P, Walliker D, Mackinnon MJ. 2001. Antimalarial drugs clear resistant parasites from partially immune hosts. Antimicrobial Agents and Chemotherapy 45:2897–2901. Davies TA, Pankuch GA, Dewasse BE, Jacobs MR, Applebaum PC. 1999. In vitro development of resistance to five quinolones and amoxicillin-clavulanate in Streptococcus pneumoniae. Antimicrobial Agents and Chemotherapy43:1177–1182. De D, Krogstad FM, Cogswell FB, Krogstad DJ. 1996. Aminoquinolines that circumvent resistance in Plasmodium falciparum in vitro. American Journal of Tropical Medicine and Hygiene55:579–583. de Lencastre H, Wu SW, Pinho MG, Ludovice AM, Filipe SR, Gardete S, Sobral R, Gill S, Chung M, Tomasz A. 1999. Antibiotic resistance as a stress response: complete sequence of a large number of chromosomal loci in Staphylococcus aureus strain COL that impact on the expression of resistance to methicillin. Microbial Drug Resistance5:163–175. de Silva NR, Chan MS, Bundy DA. 1997. Morbidity and mortality due to ascariasis: re-estimation and sensitivity analysis of global numbers at risk. Tropical Medicine and International Health2:519–528. Dixon TC, Meselson M, Guillemin J, Hanna PC. 1999. Anthrax. New England Journal of Medicine341:815–826. Djimdé A, Doumbo OK, Cortese JF, Kayentao K, Doumbo S, Diourté Y, Dicko A, Su X, Nomura T, Fidock DA, Wellems TE, Plowe CV. 2001. A molecular marker for chloroquine-resistant falciparum malaria. New England Journal of Medicine344:257–263. Doenhoff M, Kimani G, Cioli D. 2000. Praziquantel and the control of schistosomiasis. Parasitology Today16:364–366. Dorn A, Vippagunta SR, Matile H, Jaquet C, Vennerstrom JL, Ridley RG. 1998. An assessment of drug-haematin binding as a mechanism for inhibition of haematin polymerisation by quinoline antimalarials. Biochemical Pharmacology55:727–736. Dorsey G, Kamya MR, Singh A, Rosenthal PJ. 2001a. Polymorphisms in the Plasmodium falciparum pfcrt and pfmdr-1 genes and clinical response to chloroquine in Kampala, Uganda. Journal of Infectious Diseases183:1417–1420.

OCR for page 44
Dorsey G, Fidock DA, Wellems TE, Rosenthal PJ. 2001b. Mechanisms of quinoline resistance. In: Rosenthal PJ, ed. Antimalarial Chemotherapy: Mechanisms of Action, Resis tance, and New Directions in Drug Discovery. Totowa, NJ: Humana Press. Pp. 153– 172. Elliott TB, Brook I, Harding RA, Bouhaouala SS, Shoemaker MO, Knudson GB. 2002. Antimicrobial therapy for Bacillus anthracis-induced polymicrobial infection in 60Co-?-irradiated mice. Antimicrobial Agents and Chemotherapy46:3463–3471. Elliott TB, Brook I, Stiefel SM. 1990. Quantitative study of wound infection in irradiated mice. International Journal of Radiation Biology58:341–350. Fallon PG, Mubarak JS, Fookes RE, Niang M, Butterworth AE, Sturrock RF, Doenhoff MJ. 1997. Schistosoma mansoni: maturation rate and drug susceptibility of different geographic isolates. Experimental Parasitology86:29–36. Farr BM and Jarvis WR. 2002. Would active surveillance cultures help control health care-related methicillin-resistant Staphylococcus aureus infections?Infection Control and Hospital Epidemiology23:65–68. Fidock DA, Nomura T, Talley AK, Cooper RA, Dzekunov SM, Ferdig MT, Ursos LM, Sidhu AB, Naudé B, Deitsch KW, Su X, Wootton JC, Roepe PD, Wellems TE. 2000. Mutations in the P. falciparum digestive vacuole transmembrane protein PfCRT and evidence for their role in chloroquine resistance. Molecular Cell6:861–871. Filipe SR and Tomasz A.2000. Inhibition of the expression of penicillin resistance in Strepto coccus pneumoniae by inactivation of cell wall muropeptide branching genes. Proceed ings of the National Academy of Sciences97:4891–4896. Greenberg AE, Ntumbanzondo M, Ntula N, Mawa L, Howell J, Davachi F. 1989. Hospital-based surveillance of malaria-related paediatric morbidity and mortality in Kinshasa, Zaire. Bulletin of the World Health Organization67:189–196. Gryseels B, Mbaye A, De Vlas SJ, Stelma FF, Guisse F, Van Lieshout L, Faye D, Diop M, Ly A, Tchuem-Tchuente LA, Engels D, Polman K. 2001. Are poor responses to praziquantel for the treatment of Schistosoma mansoni infections in Senegal due to resistance? An overview of the evidence. Tropical Medicine and International Health6:864–873. Hansman D, Devitt L, Miles H, Riley I. 1974. Pneumococci relatively insensitive to penicillin in Australia and New Guinea. Medical Journal of Australia2:353–356. Hawley SR, Bray PG, O’Neill PM, Naisbitt DJ, Park BK, Ward SA. 1996. Manipulation of the N-alkyl substituent in amodiaquine to overcome the verapamil-sensitive chloroquine resistance component. Antimicrobial Agents and Chemotherapy40:2345–2349. Hellgren U, Kihamia CM, Mahikwano LF, Bjorkman A, Eriksson O, Rombo L. 1989. Response of Plasmodium falciparum to chloroquine treatment: relation to whole blood concentrations of chloroquine and desethylchloroquine. Bulletin of the World Health Organization67:197–202. Inglesby TV, Henderson DA, Bartlett JG, Ascher MS, Eitzen EM, Friedlander AM, Hauer J, McDade J, Osterholm MT, O’Toole T, Parker G, Perl TM, Russell PK, Tonat K. 1999. Anthrax as a biological weapon: medical and public health management. Journal of the American Medical Association281:1735–1745. Jevons MP. 1961. “Celebenin”-resistant staphylococci. British Medical Journal1:124–125. King CH. 2001. Epidemiology of schistosomiasis: determinants of transmission of infection. In: Mahmoud AAF, ed. Schistosomiasis. London: Imperial College Press. Pp. 115–132. King CH and Mahmoud AA. 1989. Drugs five years later: praziquantel. Annals of Internal Medicine110:290–296. King CH, Muchiri EM, Ouma JH. 2000. Evidence against rapid emergence of praziquantel resistance in Schistosoma haematobium, Kenya. Emerging Infectious Diseases6:585– 594.

OCR for page 44
Maguire JD, Susanti AI, Krisin, Sismadi P, Fryauff DJ, Baird JK. 2001. The T76 mutation in the pfcrt gene of Plasmodium falciparum and clinical chloroquine resistance phenotypes in Papua, Indonesia. Annals of Tropical Medicine and Parasitology95:559–572. Martin SK, Oduola AM, Milhous WK. 1987. Reversal of chloroquine resistance in Plasmo dium falciparum by verapamil. Science235:899–901. Mayor AG, Gómez-Olivé X, Aponte JJ, Casimiro S, Mabunda S, Martinho D, Barreto A, Alonso PL. 2001. Prevalence of the K76T mutation in the putative Plasmodium falciparum chloroquine resistance transporter (pfcrt) gene, and its relation to chloroquine resistance in Mozambique. Journal of Infectious Diseases183:1413–1416. Nomura T, Carlton JM, Baird JK, del Portillo HA, Fryauff DJ, Rathore D, Fidock DA, Su X, Collins WE, McCutchan TF, Wootton JC, Wellems TE. 2001. Evidence for different mechanisms of chloroquine resistance in two Plasmodium species that cause human malaria. Journal of Infectious Diseases183:1653–1661. Oliveira DC, Tomasz A, de Lencastre H. 2002. The secrets of success of a human pathogen: molecular evolution of pandemic clones of methicillin-resistant Staphylococcus aureus. Lancet Infectious Diseases2:180–189. Pagola S, Stephens PW, Bohle DS, Kosar AD, Madsen SK. 2000. The structure of malaria pigment ß-haematin. Nature404:307–310. Partnership for Child Development. 1997. Better health, nutrition, and education for the school-aged child. The Partnership for Child Development. Transactions of the Royal Society for Tropical Medicine and Hygiene91:1–2. Payne D. 1987. Spread of chloroquine resistance in Plasmodium falciparum. Parasitology Today3:241–246. Peters W. 1987. Resistance in human malaria IV: 4-aminoquinolines and multiple resistance. In: Chemotherapy and Drug Resistance in Malaria. London: Academic Press. Pp. 659– 786. Peters W. 1989. Changing pattern of antimalarial drug resistance. Journal of the Royal Soci ety of Medicine82 Suppl 17:14–17. Peterson VM, Adamovicz JJ, Elliott TB, Moore MM, Madonna GS, Jackson WE 3rd, Ledney GD, Gause WC. 1994. Gene expression of hematoregulatory cytokines is elevated endogenously following sublethal ?-irradiation and is differentially enhanced by therapeutic administration of biological response modifiers. Journal of Immunology153:2321– 2330. Pillai DR, Labbe AC, Vanisaveth V, Hongvangthong B, Pomphida S, Inkathone S, Zhong K, Kain KC. 2001. Plasmodium falciparum malaria in Laos: chloroquine treatment outcome and predictive value of molecular markers. Journal of Infectious Diseases183:789– 795. Pittet D. 2002. Promotion of hand hygiene: magic, hype, or scientific challenge?Infection Control and Hospital Epidemiology23:118–119. Rasoanaivo P, Ratsimamanga-Urverg S, Frappier F. 1996. Reversing agents in the treatment of drug-resistant malaria. Current Medicinal Chemistry3:1–10. Reed MB, Saliba KJ, Caruana SR, Kirk K, Cowman AF. 2000. Pgh1 modulates sensitivity and resistance to multiple antimalarials in Plasmodium falciparum. Nature403:906–909. Renganathan E and Cioli D.1998. An international initiative on praziquantel use. Parasitol ogy Today14:390–391. Ridley RG, Hofheinz W, Matile H, Jaquet C, Dorn A, Masciadri R, Jolidon S, Richter WF, Guenzi A, Girometta MA, Urwyler H, Huber W, Thaithong S, Peters W. 1996. 4-aminoquinoline analogs of chloroquine with shortened side chains retain activity against chloroquine-resistant Plasmodium falciparum. Antimicrobial Agents and Chemotherapy 40:1846–1854.

OCR for page 44
Rieckmann KH, Davis DR, Hutton DC. 1989. Plasmodium vivax resistant to chloroquine? Lancet2:1183–1184. Sá-Leão R, Santos Sanches I, Couto I, Alves CR, de Lencastre H. 2001. Low prevalence of methicillin-resistant strains among Staphylococcus aureus colonizing young and healthy members of the community in Portugal. Microbial Drug Resistance7:237–241. Sá-Leão R, Tomasz A, Santos Sanches I, Brito-Avô A, Vilhelmsson SE, Kristinsson KG, de Lencastre H. 2000. Carriage of internationally spread epidemic clones of Streptococcus pneumoniae with unusual drug resistance patterns in children attending day care centers in Lisbon, Portugal . The Journal of Infectious Diseases182:1153–1160. Sowunmi A, Oduola AM, Ogundahunsi OA, Falade CO, Gbotosho GO, Salako LA. 1997. Enhanced efficacy of chloroquine-chlorpheniramine combination in acute uncomplicated falciparum malaria in children. Transactions of the Royal Society of Tropical Medicine and Hygiene91:63–67. Sturrock RF. 2001. The schistosomes and their intermediate hosts. In: Mahmoud AAF, ed. Schistosomiasis. London: Imperial College Press. Pp. 7–83. Sullivan DJ, Gluzman IY, Russell DG, Goldberg DE. 1996. On the molecular mechanism of chloroquine’s antimalarial action. Proceedings of the National Academy of Sciences 93:11865–11870. Tomasz A. 1994. Multiple-antibiotic-resistant pathogenic bacteria. A report on the Rockefeller University workshop. New England Journal of Medicine330:1247–1251. Tomasz A. 2000. Lessons from the first antibiotic era. In: Andrew PW, Oyston P, Smith GL, Stewart-Tull DE, eds. Fighting Infection in the 21st Century. Society for General Microbiology Millenium Meeting Symposium Volume. Oxford, UK: Blackwell Science. Pp. 198–216. Trape JF, Pison G, Preziosi MP, Enel C, Desgrees du Lou A, Delaunay V, Samb B, Lagarde E, Molez JF, Simondon F. 1998. Impact of chloroquine resistance on malaria mortality. Comptes Rendus de l’Académie des Sciences. Série III, Sciences de la Vie321:689–697. Van Wyk JA. 2001. Refugia—overlooked as perhaps the most potent factor concerning the development of anthelmintic resistance. Onderstepoort Journal of Veterinary Research 68:55–67. Verdier F, Le Bras J, Clavier F, Hatin I, Blayo MC. 1985. Chloroquine uptake by Plasmo dium falciparum-infected human erythrocytes during in vitro culture and its relationship to chloroquine resistance. Antimicrobial Agents and Chemotherapy27:561–564. Vieira PP, das Gracas Alecrim M, da Silva LH, González-Jiménez I, Zalis MG. 2001. Analysis of the PfCRT K76T mutation in Plasmodium falciparum isolates from the Amazon region of Brazil. Journal of Infectious Diseases183:1832–1833. Vippagunta SR, Dorn A, Matile H, Bhattacharjee AK, Karle JM, Ellis WY, Ridley RG, Vennerstrom JL. 1999. Structural specificity of chloroquine-hematin binding related to inhibition of hematin polymerization and parasite growth. Journal of Medicinal Chemis try42:4630–4639. Warren KS. 1982. Selective primary health care: strategies for control of disease in the developing world. I. Schistosomiasis. Reviews of Infectious Diseases4:715–726. Wellems TE and Plowe CV. 2001. Chloroquine-resistant malaria. Journal of Infectious Dis eases184:770–776. Whitby M. 1997. Drug resistant Plasmodium vivax malaria. Journal of Antimicrobial Che motherapy40:749–752. WHO (World Health Organization). 1993. The Control of Schistosomiasis: Second Report of the WHO Expert Committee.Geneva: WHO. WHO. 2000. Overcoming Antimicrobial Resistance: WHO Report on Infectious Diseases 2000. [Online]. Available: http://www.who.int/infectious-disease-report/2000/.

OCR for page 44
William S, Sabra A, Ramzy F, Mousa M, Demerdash Z, Bennett JL, Day TA, Botros S. 2001. Stability and reproductive fitness of Schistosoma mansoni isolates with decreased sensitivity to praziquantel. International Journal of Parasitology31:1093–1100. Wootton JC, Feng X, Ferdig MT, Cooper RA, Mu J, Baruch DI, Magill AJ, Su XZ. 2002. Genetic diversity and chloroquine selective sweeps in Plasmodium falciparum. Nature 418:320–323.