M E T H O D S I N M O L E C U L A R M E D I C I N E TM Edited by Denise L. Doolan Malaria Methods and Protocols Humana Press Humana Press Edited by Denise L. Doolan Malaria Methods and Protocols Vector Incrimination and EIR 3 3 From: Methods in Molecular Medicine, Vol. 72: Malaria Methods and Protocols Edited by: Denise L. Doolan © Humana Press, Inc., Totowa, NJ 1 Vector Incrimination and Entomological Inoculation Rates John C. Beier 1. Introduction This chapter provides standard methods for the incrimination of Anopheles mos- quito species serving as malaria vectors and associated methods for measuring the intensity of transmission. In any malaria-endemic area, one or more species of Anoph- eles mosquitoes serve as malaria vectors. To show that an Anopheles mosquito species serves as a malaria vector in nature, it is necessary to demonstrate: 1. An association in time and space between the Anopheles species of mosquito and cases of malaria in humans. After study sites are selected, longitudinal field studies are established to sample mosquito populations. Adult mosquitoes are sampled by using trapping tech- niques such as landing/biting collections, light traps, pyrethrum spray catches inside houses, and outdoor aspiration collections. Larval mosquitoes developing in aquatic habi- tats normally are sampled by dipping methods. Mosquitoes are identified by standard taxo- nomic methods and also by molecular methods if mosquitoes belong to a species complex. The standard methods for performing landing/biting collections are described in this chap- ter; other types of mosquito trapping methods are described in refs. 1 and 2. 2. Evidence of direct contact between the Anopheles species and humans. Catching a mos- quito biting humans through landing/biting catches conclusively establishes contact between that mosquito species and humans. A second method involves immunologically identifying human blood in the abdomen of field-captured Anopheles mosquitoes. A direct enzyme-linked immunosorbent assay (ELISA) suitable for bloodmeal identification of African malaria vectors is described in this chapter (3). 3. Evidence that the Anopheles species harbors malaria sporozoites in the salivary glands. Sporozoites may be detected in mosquitoes through the dissection and microscopic examination of mosquito salivary glands (4) or through ELISA methods (5). Both methods are described in this chapter. The process of vector incrimination is often done in conjunction with longitudinal field studies to measure the intensity of transmission. In an endemic area, the intensity of transmission is determined by calculating the entomological inoculation rate (EIR), which is the product of the mosquito biting rate times the proportion of mosquitoes with sporozoites. EIRs, calculated as the sum total for each individual vector species of mosquito, are expressed in terms of average numbers of infective bites per person per unit time. For example, EIRs in endemic areas of Africa generally range from 1 to >1000 infective bites per year (6). In this chapter, methods are described for calculat- 4 Beier ing EIRs from landing/biting catches and determinations of sporozoite rates. Further details and references are provided on the use of EIRs for epidemiological studies and for determining levels of control necessary for achieving reductions in malaria preva- lence and the incidence of severe disease. 2. Materials 2.1. Equipment 1. Mouth aspirators for collecting mosquitoes. 2. Flashlights. 3. Hand-held global position satellite (GPS) system receiver. 4. Low-intensity kerosene lantern. 5. Screened paper pint cups for holding live mosquitoes. 6. Labels and/or permanent marker pens. 7. Glass microscope slides. 8. Phase-contrast compound microscope and dissecting microscope with light source. 9. Surgical scalpel blades. 10. Glass rods or plastic pestles for grinding mosquitoes. 11. 1.8-mL plastic tubes with snap-on caps for holding mosquito samples. 12. 15- and 50-mL tubes for mixing ELISA reagents. 13. Freezer for storing mosquito samples at –20 or –70°C. 14. Refrigerator. 15. 8-Channel manifold attached to 60-mL plastic syringe. 16. Polyvinyl chloride (PVC) microtiter plates. 17. Absorbent tissue paper. 18. ELISA plate reader. 2.2. Reagents 1. Chloroform, ether, and/or 70% ethanol for killing mosquitoes. 2. Physiological saline or medium-199 (M-199) for dissecting mosquitoes. 3. Phosphate-buffered saline (PBS). 4. PBS–Tween-20 (PBS–Tw20) wash solution for ELISA: Add 500 µL of Tween-20 to 1 L of PBS, mix, and store in a refrigerator. 5. Boiled casein blocking buffer (BB) for ELISA: Suspend 5.0 g casein in 100 mL of 0.1 N sodium hydroxide and bring to a boil while stirring on a hot plate. After casein has dis- solved, slowly add 900 mL of PBS, allow to cool, and adjust the pH to 7.4 with hydrochlo- ric acid (HCl). Add 0.1 g thimerosal and 0.02 g phenol red. Mix well using a magnetic stirrer and store in a refrigerator; shelf life is 7 to 10 d. 6. Blocking buffer Nonidet P-40 (BB–NP-40) for grinding mosquitoes: prepare by adding 5 µL of NP-40 to each 100 µL BB and mixing. Make fresh daily. 7. Capture and conjugated monoclonal antibodies for sporozoite ELISA tests. 8. Peroxidase substrate (ABTS) and phosphatase substrate. 9. Recombinant proteins as positive controls for sporozoite ELISA tests. 10. Host-specific peroxidase conjugates (anti-host IgG, H&L) and phosphatase-labeled anti- bovine IgG (H&L) for bloodmeal ELISA. 11. Host sera as controls for the bloodmeal ELISA. 3. Methods 3.1. Site Selection and Mosquito Sampling Stations 1. Study sites are selected based on study objectives that may be related to epidemiological studies, malaria control operations, vaccine or drug testing under natural conditions, or an Vector Incrimination and EIR 5 abundance of vector species of mosquitoes. Study sites may range in size from a cluster of a few houses to whole communities. Prior to selecting and working in sites, it is advisable to discuss study objectives and operations with community leaders and residents and to obtain their consent for the field studies (see Note 1). 2. Normally, it is necessary to develop study site maps based on either traditional mapping methods or through geographic information systems (GIS) using GPS receivers for deter- mining latitudes and longitudes of houses and other landmark features within sites. The maps are used to facilitate field studies logistically and serve as a foundation for the analy- sis of spatial data on mosquito populations and transmission. 3. Sampling stations for mosquito trapping are selected within sites. For highly endophilic and anthropophilic mosquitoes like the African malaria vectors, sampling stations are nor- mally houses or homesteads comprising family units of houses. For exophilic or zoophilic Anopheles species, sampling stations can be either outdoor areas or animal sheds. The number of sampling stations depends upon logistical capabilities such as the number of mosquito collectors available and the expected frequency of sampling within the study sites. Sampling stations are normally fixed and used repeatedly throughout the duration of field studies. Alternatively, sampling stations can be selected randomly during each sam- pling period. While this is sometimes useful from a statistical perspective, we have found that this approach makes it more difficult to obtain good cooperation from communities. 3.2. Landing/Biting Catches of Anopheles Mosquitoes Landing/biting catches of Anopheles mosquitoes on human volunteers (see Note 2) are performed either by individual human collectors, by pairs of collectors, or by up to four collectors working simultaneously. Collections are normally performed at night, during the biting cycles of the Anopheles mosquitoes. Each collector is responsible for catching, by mouth aspirator with the aid of a flashlight, mosquitoes that are attracted to and in the process of biting humans (i.e., host-seeking mosquitoes). The trapping technique simulates the natural situation whereby mosquitoes contact and bite humans, and so it is regarded as the gold standard. For each malaria vector field study, it is necessary to evaluate all other trapping methods for evaluating host contact against the gold standard. Procedurally, landing/biting catches are performed as follows: 1. Collectors with their arms and legs exposed are seated in chairs or on mats on the ground at sampling stations. It is common to perform indoor and outdoor biting catches simulta- neously at the same sampling stations, with the outdoor collectors positioned at least 5 m from surrounding houses. Trapping inside houses provides information on the numbers of mosquitoes biting inside, while performing the sampling outdoors provides comparable information on outdoor biting rates. 2. Collectors catch landing/biting mosquitoes from themselves and from their partners with a hand-held mouth aspirator (or mechanical aspirator). Each collector uses a flashlight to locate landing/biting mosquitoes. Additional background light from a low-intensity lan- tern is advisable. 3. Each aspirated mosquito is placed in screened pint cups, labeled according to sampling station. Some studies also segregate mosquito collections by hour of capture, and this requires additional cups labeled by hour. 4. Landing/biting collections are normally performed throughout the night as dictated by the natural biting habits of the target mosquitoes. Logistically, it is feasible for individuals or teams of collectors to work one-half hour every hour throughout the night. Alternatively, it is feasible for half the team to work continuously during the first half of the night and the rest of the team to work the second half of the night. 6 Beier 5. After collections, mosquitoes in cups are normally killed either by freezing or by exposure to chloroform or ether. For immediate processing, mosquitoes may be aspirated out of cups and blown into 70% ethanol followed by transfer to PBS or M-199. Mosquitoes may also be stored in Carnoy’s solution for cytogenetic studies or for longer-term storage before processing (see Chapter 8). 6. Mosquitoes are identified according to taxonomic methods or by molecular techniques (see Chapter 8). 7. The biting rate for each mosquito species is calculated as the number of mosquitoes per person per unit of sampling effort. For example, a biting rate of 2 per day is derived from one collector who catches one mosquito while working throughout the night in half-hour shifts. A biting rate of 40 per day is derived from a team of two collectors working in half- hour shifts during the whole night and catching 40 mosquitoes. 3.3. Determination of Sporozoite Rates in Anopheles Mosquitoes 3.3.1. Dissection and Microscopic Examination of Mosquito Salivary Glands 1. Salivary gland dissections are performed on mosquitoes freshly killed by freezing, expo- sure to ether, or by blowing into 70% ethanol. 2. Place an individual mosquito on a glass slide, with head in contact with a small drop of physiological saline or PBS or M-199. 3. View slide containing mosquito on a dissecting microscope at ×10 to ×30. 4. Hold two dissecting needles, which can be made conveniently by placing the 27-gage needle from a 1-mL tuberculin syringe on the end of the movable shaft (rubber stopper removed), between your thumb and forefinger. Place one needle (bevel down) on the tho- rax of the mosquito while placing the other needle against the mosquito head (bevel facing toward the head). Simultaneously place pressure on the thorax while pulling the head away from the thorax. As the head moves away from the thorax, observe the salivary glands and cut them with the needle controlling the head. The cut should be made in one continuous motion as soon as the glands are seen; otherwise, it is necessary to reposition the head and try again. Sometimes the glands become stuck in the mosquito thorax, and it is necessary to tease apart the tissue to locate and cut the glands. 5. After severing the salivary glands, remove the head and thorax and any other extraneous tissue. 6. Place a glass cover slip over the salivary glands, now lying in the dissection media. 7. Transfer the slide to a compound microscope and observe the preparation at ×100 to locate the salivary glands. 8. Apply gentle pressure to the cover slip to disrupt the glands and then search at ×400 the entire area of the salivary glands for sporozoites (which normally measure about 1 × 10 µm). Experienced dissectors can typically dissect a mosquito within 1 min and reliably exam- ine the preparation within 2 min. 9. Sporozoite infections are normally scored according to the number of sporozoites observed: 1+ (1–10 sporozoites), 2+ (11–100 sporozoites), 3+ (101–1000), and 4+ (>1000 sporozoites). 10. Record results. Normally, each field-collected mosquito is given a unique identifier (see Note 3). 11. Standard procedures are also available for removing sporozoite material from slides and testing the sporozoites by ELISA to determine Plasmodium species (7). Various addi- tional procedures are available for determining sporozoite loads, the number of sporozoi- tes found in the salivary glands of individual mosquitoes (8,9). 3.3.2. Sporozoite ELISA Methods ELISA methods exist for testing field-collected mosquitoes for sporozoites repre- senting each of the four species of Plasmodium affecting humans (5,10,11). The sporo- Vector Incrimination and EIR 7 zoite ELISA detects circumsporozoite protein that is either from intact sporozoites or in soluble form within the mosquito. Based on comparisons with the gold standard dissection method, the sporozoite ELISA provides a reasonable estimate of the true sporozoite rate in wild-caught mosquitoes (12). 1. Prepare the mosquito sample for ELISA testing. Label sets of 1.8-mL tubes with the cor- responding mosquito sample numbers. Add 50 µL of BB–NP-40 to each vial. Using a sharp clean surgical blade, cut the mosquito between the thorax and the abdomen (nor- mally done on a filter paper). Transfer the head–thorax with forceps to the labeled tube, and transfer the abdomen to the corresponding tube for bloodmeal identification if the mosquito is blood-fed. If the mosquito is not blood-fed or no bloodmeal analysis is required, discard abdomen. Grind the mosquito in the tube using a nonabsorbent glass rod or plastic pestle. Add 200 µL of the BB to bring the total sample volume to 250 µL. To avoid contamination, clean the pestle and wipe it dry before grinding the next sample. Repeat the procedure until all samples are prepared. Arrange samples in numbered order within storage boxes and keep samples in a freezer at –20 or –70°C until testing. 2. Coat number-coded ELISA plates with monoclonal antibody (MAb). In each well, add 50 µL of the diluted capture MAb. Cover the plates with another clean ELISA plate and incubate for 30 min at room temperature in subdued light. 3. Block the plates. Using an 8-channel manifold attached to a vacuum pump, aspirate the capture MAb from the microtiter plate. Bang the plate hard on an absorbent tissue paper or gauze to ensure complete dryness. Fill each well with BB using a manifold attached to a 60-mL syringe. Incubate for 1 h at room temperature in subdued light. 4. Load the plates with mosquito samples. Aspirate the blocking buffer from the wells using the manifold attached to a vacuum pump and bang plate to complete dryness. Place 50 µL of 100, 50, 25, 12, 6, 3, 1.5, 0 pg of positive control recombinant protein in the first column wells. Into the second column, add 50 µL per well of the negative controls; nor- mally, field-collected male Anopheles mosquitoes or culicine mosquitoes are used as nega- tive controls. Load 50 µL of each mosquito sample to the remaining wells of the plate, checking carefully that numbered mosquito samples are placed in the wells according to the completed ELISA data form. Cover the plate and incubate for 2 h at room temperature in subdued light. 5. Add peroxidase-conjugated monoclonal antibody. After 2 h, aspirate the triturate from the wells and wash the plate two times with PBS-Tw20. Add 50 µL of the peroxidase-labeled enzyme and incubate for 1 h at room temperature. 6. Add the substrate. Aspirate the enzyme conjugate from the wells and wash three times with PBS-Twn 20. Using a multichannel pipet, add 100 µL of ABTS substrate and incu- bate for 30 min. Positive reactions, which appear green, can be determined by reading plates at 414 nm using an ELISA plate reader; absorbance values two times the mean of negative controls provides a valid cutoff for sample positivity (13). Alternatively, results can be read visually with a high degree of accuracy (14). Record results for each tested mosquito. 3.4. Bloodmeal ELISA Methods Both direct and indirect ELISA procedures are routinely used to identify bloodmeals of wild-caught mosquitoes. Strategies for using ELISAs for bloodmeals depend upon study objectives. For example, Edrissian et al. (15) used a direct ELISA to screen over 5000 Anopheles for human blood; they reported that an experienced technician could easily screen over 1000 samples per week. Burkot and DeFoliart (16) used an indirect ELISA to identify 16 host sources, including wild animals. Their studies involved pro- 8 Beier ducing antisera for each host tested. To bypass extensive production of antisera and to shorten the overall testing time, we developed a simple direct ELISA that uses only commercially available reagents (3). The test reliably detects bloodmeals from humans and from a spectrum of domestic animals for which conjugated antisera are commer- cially available. The test employs a two-step screening for human and cow bloodmeals, making it particularly useful in Africa where major malaria vectors feed primarily on humans and cows. The assay is performed as follows: 1. Prepare wild-caught half-gravid to freshly fed mosquitoes by cutting them transversely at the thorax between the first and third pairs of legs (under a dissecting microscope, ×10– 20). In a labeled tube, place the posterior part of the mosquito containing the bloodmeal in 50 µL PBS and grind with a pestle or pipet repeatedly. Dilute sample 1:50 with PBS and freeze samples at –20°C until testing. 2. Load 96-well polyvinyl microtiter plates with mosquito bloodmeal samples by adding 50 µL of each sample per well. On the same plate, add 50-µL samples of positive control antis- era for human and cow (diluted 1:500 in PBS), and four or more negative control unfed female mosquitoes or male mosquitoes obtained from the same field collections and handled as above. Cover and incubate at room temperature for 3 h (or overnight). 3. Wash each well twice with PBS-Tw20. 4. Add 50 µL of host-specific conjugate (anti-host IgG, H&L) diluted 1:2,000 (or as deter- mined in control tests) in 0.5% BB containing 0.025% Tween-20, and incubate 1 h at room temperature. 5. Wash wells three times with PBS–Tw-20. 6. Add 100 µL of ABTS peroxidase substrate to each well. 7. After 30 min, read each well with an ELISA reader. Samples are considered positive if absorbance values exceed the mean plus three standard deviations of four negative con- trol, unfed female, or male mosquitoes. The dark green positive reactions for peroxidase (or the dark yellow reactions for phosphatase) may also be determined visually (14). 8. The following modification is used in the two-step procedure for determining a second host source in the same microtiter plate well where mosquito samples were screened for human blood (3). A second conjugate, phosphatase-labeled anti-bovine IgG (1:250 dilu- tion of a 0.5 mg/mL stock solution) is added to the peroxidase-labeled anti-human IgG solu- tion (step 4). Screen bloodmeals first for human IgG by adding peroxidase substrate, and after reading absorbance at 30 min, wash the wells three times with PBS–Tw20. Add 100 µL of phosphatase substrate and read plates after 1 h to determine positive cow reactions. 9. Each laboratory should initially establish the sensitivity and specificity of the assay for each conjugated antisera and different lots of reagents. The assay can detect sera diluted to around 1:10,000,000. The degree to which commercially available antisera crossreact with sera from different hosts varies according to manufacturers. For standardization and to reduce levels of nonspecific reactivity, it is sometimes necessary to add 1:500 dilutions of heterologous sera to the conjugate solutions (see step 4 ). It is important to note that the assay works equally well with frozen, dried, or Carnoy’s fixed mosquito samples, and that each 1-mg vial of conjugated antisera can be aliquoted, frozen, and used to test up to 20,000 mosquito samples. 3.5. Entomological Inoculation Rates ( see Note 4) 3.5.1. Calculation of EIRs The EIR is calculated as the product of the mosquito biting rate and the sporozoite rate. Table 1 provides an example for the calculation of the EIR from site-specific data on mosquito biting rates and sporozoite rates. Beyond calculating daily EIRs, it is also Vector Incrimination and EIR 9 useful to calculate monthly or annual EIRs based on averaged values of biting rates and sporozoite rates. For the example given in Table 1, if these were the only sample data available, then an estimate of the monthly EIR could be obtained by multiplying the daily EIR of 0.60 by 30 d to yield an estimated monthly EIR of 18. For field studies, it is advisable to have two or more point determinations of biting rates per month and statistically reliable estimates of sporozoite rates. 3.5.2. Relationships Between EIRs and Measures of Human Malaria Time series data on EIRs for given sites can be related directly to measures of human malaria in several simple ways. Graphically, measures of the EIR and human malaria such as prevalence or incidence can be graphed along the y-axis and related to sam- pling time points on the x-axis. In addition, such temporal changes in EIR and infection or disease can be related to environmental parameters such as temperature and rainfall. The same data can be graphed with EIRs on the x-axis and human malaria data on the y- axis. This approach, when combined with regression analysis, provides both a graphical and a statistical account of the variation in human infection or disease explained by EIRs. Several studies in Africa provide good examples of how EIRs can be related to the following: 1. Incidence of P. falciparum infection in children (17). 2. Incidence of severe life-threatening cases of P. falciparum in children (18,19). 3. Prevalence of P. falciparum (6). It is important to note that malaria prevalence data, which is often used as the basis for guiding control operations, is not a sensitive indicator of the intensity of malaria transmission by vector populations. Malaria prevalence rates from 40 to >90% can occur at any EIR exceeding one infective bite per person per year. Control operations therefore need to be guided by both entomological data on EIRs and traditional mea- sures of human malaria infection and disease. 4. Notes 1. Ethical concerns must be addressed for each malaria vector field study. Mosquito trapping in malaria endemic areas normally involves local mosquito collectors who are recruited from study communities. For landing/biting mosquito sampling methods, local mosquito Table 1 Entomological Inoculation Rate Species Biting rate a Sporozoite rate (%) b Daily EIR A10 5.00 0.50 B4 2.00 0.08 C2 1.00 0.02 Total 16 3.75 0.60 The EIR is calculated as the product of the biting rate times the sporozo- ite rate. In this hypothetical example, there are three vector species of Anopheles mosquitoes. The daily EIR of 0.60 infective bites per person per night is calculated as the sum total of EIRs for each of the three species. a Number of mosquitoes per person per night. b Percentage of mosquitoes with salivary gland sporozoites by dissection or ELISA. 10 Beier collectors normally do not face any excess risks for malaria infection beyond what they would normally experience sleeping in their own homes. Some studies have traditionally offered malaria prophylactic drugs to collectors. However, in highly endemic areas, this practice goes beyond normal protective measures of the community and may even be detrimental to the long-term health of the collectors. It is advisable that malaria field stud- ies make sure that mosquito collectors have proper access to curative antimalarial drugs and health facilities whenever they develop symptomatic malaria infections. 2. The landing/biting catch method is the gold standard for determining human contact with mosquitoes. Other methods such as pyrethrum spray catches or CDC light traps may be used to estimate rates of human biting for each Anopheles species. However, for each study area, it is necessary to establish quantitative relations between the sampling method proposed and the gold standard method. Estimates of correction factors from regression analysis based on data from comparative sampling for one site do not generally hold uni- versally throughout the ranges of mosquitoes (20). 3. Care must be taken with data management. Malaria vector studies normally yield large numbers of mosquitoes from different trapping methods that are processed by a variety of different methods ranging from taxonomic identifications to sporozoite ELISA testing. Primary data sets can be established in matrix format with each mosquito represented by rows and columns represented by collection and processing data. Prospects of having >15 variables of data for each individual mosquito and over 50,000 mosquitoes in a single dataset demand careful attention in terms of data entry, data management, and analysis. 4. The EIR is a direct measure of the intensity of malaria transmission by vector populations. In malaria endemic areas outside Africa and Papua New Guinea, annual EIRs may be lower than one infective bite per person per year (21). The low EIRs are often due to sporozoite rates substantially less than 1%. In some areas of Central and South America, for example, it is not uncommon to find fewer than 1 in 1000 mosquitoes infected with malaria parasites. Under such conditions, investigators may alternatively calculate the vec- torial capacity (VC), an indirect measure of the potential for transmission that considers the biting rate of the vector population, the human blood-feeding rate, the vector survival rate, and the extrinsic incubation period of the malaria parasite (22). Some of the practical limita- tions and errors associated with the use of the VC are discussed by Dye (23). Acknowledgments This work was supported by the National Institutes of Health grants AI29000, AI45511, and TW01142. References 1. Service, M. W. (1976) Mosquito Ecology. Wiley, New York. 2. Service, M. (1993) Mosquito Ecology: Field Sampling Methods. Elsevier Applied Science, New York. 3. Beier, J. C., Perkins, P. V., Wirtz, R. A., Koros, J., Diggs, D., Gargan, T. P., and Koech, D. K. (1988) Bloodmeal identification by direct enzyme-linked immunosorbent assay (ELISA), tested on Anopheles (Diptera: Culicidae) in Kenya. J. Med. Entomol. 25, 9–16. 4. WHO (1975) Manual on Practical Entomology in Malaria. Part II. Methods and Techniques. WHO Offset Publication 13, Geneva. 5. Wirtz, R. A. and Burkot, T. R. (1991) Detection of malaria parasites in mosquitoes. Chap 4 in Advances in Disease Vector Research, vol. 8, Springer-Verlag, New York, pp. 77–106. 6. Beier, J. C., Killeen, G. F., and Githure, J. I. (1999) Short report: Entomological inoculation rates and Plasmodium falciparum malaria prevalence in Africa. Am. J. Trop. Med. Hyg. 61, 109–113. 7. Beier, J. C., Copeland, R. S., Onyango, F. K., Asiago, C. M., Ramadhan, M., Koech, D. K., and Roberts, C. R. (1991) Plasmodium species identification by ELISA for sporozoites removed from dried dissection slides. J. Med. Entomol. 28, 533–536. 8. Kabiru, E. W., Mbogo, C. M., Muiruri, S. K., Ouma, J. H., Githure, J. I., and Beier, J. C. (1997) Vector Incrimination and EIR 11 Sporozoite loads of naturally infected Anopheles in Kilifi District, Kenya. J. Am. Mosq. Control Assoc. 13, 259–262. 9. Beier, J. C., Onyango, F. K., Ramadhan, M., Koros, J. K., Asiago, C. M., Wirtz, R. A., et al. (1991) Quantitation of malaria sporozoites in the salivary glands of wild Afrotropical Anopheles. Med. Vet. Entomol. 5, 63–70. 10. Wirtz, R. A., Burkot, T., Graves, P. M., and Andre, R. G. (1987) Field evaluation of enzyme-linked immunosorbent assays for Plasmodium falciparum and Plasmodium vivax sporozoites in mosqui- toes (Diptera: Culicidae) from Papua New Guinea. J. Med. Entomol. 24, 433–437. 11. Beier, J. C., Perkins, P. V., Wirtz, R. A., Whitmire, R. E., Mugambi, M., and Hockmeyer, W. T. (1987) Field evaluation of an enzyme-linked immunosorbent assay (ELISA) for Plasmodium falciparum sporozoite detection in anopheline mosquitoes from Kenya. Am. J. Trop. Med. Hyg. 36, 459–468. 12. Beier, J. C., Perkins, P. V., Koros, J. K., Onyango, F. K., Gargan, T. P., Wirtz, R. A., et al. (1990) Malaria sporozoite detection by dissection and ELISA to assess infectivity of Afrotropical Anoph- eles (Diptera: Culicidae). J. Med. Entomol. 27, 377–384. 13. Beier, J. C., Asiago, C. M., Onyango, F. K., and Koros, J. K. (1988) ELISA absorbance cut-off method affects malaria sporozoite rate determination in wild Afrotropical Anopheles. Med. Vet. Entomol. 2, 259–264. 14. Beier, J. C. and Koros, J. K. (1991) Visual assessment of sporozoite and bloodmeal ELISA samples in malaria field studies. J. Med. Entomol. 28, 805–808. 15. Edrissian, G. H., Manouchehry, A. V., and Hafizi, A. (1985) Application of an enzyme-linked immunosorbent assay (ELISA) for determination of the human blood index in anopheline mosqui- toes collected in Iran. J. Am. Mosq. Control Assoc. 1, 349–352. 16. Burkot, T. R. and DeFoliart, G. R. (1982) Bloodmeal sources of Aedes triseriatus and Aedes vexans in a southern Wisconsin forest endemic for LaCrosse encephalitis virus. Am. J. Trop. Med. Hyg. 31, 376–381. 17. Beier, J. C., Oster, C. N., Onyango, F. K., Bales, J. D., Sherwood, J. A., Perkins, P. V., et al. (1994) Plasmodium falciparum incidence relative to entomologic inoculation rates at a site proposed for testing malaria vaccines in western Kenya. Am. J. Trop. Med. Hyg. 50, 529–536. 18. Mbogo, C. N., Snow, R. W., Kabiru, E. W., Ouma, J. H., Githure, J. I., Marsh, K., and Beier, J. C. (1993) Low-level Plasmodium falciparum transmission and the incidence of severe malaria infec- tions on the Kenyan coast. Am. J. Trop. Med. Hyg. 49, 245–253. 19. Mbogo, C. N., Snow, R. W., Khamala, C. P., Kabiru, E. W., Ouma, J. H., Githure, J. I., et al. (1995) Relationships between Plasmodium falciparum transmission by vector populations and the inci- dence of severe disease at nine sites on the Kenyan coast. Am. J. Trop. Med. Hyg. 52, 201–206. 20. Mbogo, C. N., Glass, G. E., Forster, D., Kabiru, E. W., Githure, J. I., Ouma, J. H., and Beier, J. C. (1993) Evaluation of light traps for sampling anopheline mosquitoes in Kilifi, Kenya. J. Am. Mosq. Control Assoc. 9, 260–263. 21. Alles, H. K., Mendis, K. N., and Carter, R. (1998) Malaria mortality rates in South Asia and in Africa: implications for malaria control. Parasitol. Today 14, 369-375. 22. Garrett-Jones, C. and Shidrawi, G. R. (1969) Malaria vectorial capacity of a population of Anoph- eles gambiae. Bull. World Health Organ. 40, 531–545. 23. Dye, C. (1986) Vectorial capacity: must we measure all its components? Parasitol. Today 2, 203–209. [...]... Parasite density and malaria morbidity in the Pakistani Punjab Am J Trop Med Hyg 61, 791–801 18 Smith, T., Schellenberg, J A., and Hayes, R (1994) Attributable fraction estimates and case definitions for malaria in endemic areas Stat Med 13, 2345–2358 19 McGuinness, D., Koram, K., Bennett, S., Wagner, G., Nkrumah, F., and Riley, E (1998) Clinical case definitions for malaria: clinical malaria associated... and the transmission of malaria Within ranges of temperature (20–30°C) and humidity (>60%) that vary for each vector species, the mosquito survives and is capable of transmitting malaria When the limits of temperature and humidity tolerance are exceeded, the vectors die, and the risk of malaria evaporates Variations of temperature and humidity within the viable range for mosquitoes can also affect the... setting where malaria is reported on the basis of microscopically confirmed infection, and the proportion of fevers caused by malaria is relatively low, the PCD rate may serve as a reliable estimate of risk of malaria relative to other febrile illnesses in the community In general, this scenario is true where malaria is hypo- to mesoendemic and health-care providers are more likely to report malaria on... provinces, and nations The means of deriving the numerator for the API, “cases of malaria , varies a great deal Comparisons of risk based on the API demand consideration of the sources and case definitions; for example, clinical diagnosis versus smear-confirmed diagnosis for reported total cases The API numerator often represents a hybrid of PCD and active surveillance methods (see below), and the relative... U., and Meyer, C G (1999) High rate of mixed and subpatent malaria infections in Southwest Nigeria Am J Trop Med Hyg 61, 339–343 3 Pribadi, W., Sutanto, I., Atmosoedjono, S., Rasidi, R., Surya, L K., and Susanto, L (1998) Malaria situation in several villages around Timika South Central Irian Jaya, Indonesia Southeast Asian J Trop Med Public Health 29, 228–235 4 Hii, J., Dyke, T., Dagoro, H., and Sanders,... Estimates of Risk of Malaria 2.1 Environmental (Rainfall, Altitude, Temperature) Transmission of malaria requires mosquito vectors in the genus Anopheles These insects exhibit exquisite sensitivity to the environmental parameters of temperature and humidity Thus, rainfall, altitude, and temperature govern the activity and abundance of anopheline mosquitoes and the transmission of malaria Within ranges... beta-Arteether against malaria parasites in vitro and in vivo Am J Trop Med Hyg 48, 377–384 12 Sinden, R E (1996) Infection of mosquitoes with rodent malaria, in Molecular Biology of Insect Disease Vectors: A Methods Manual (Crampton, J M., Beard, C B., and Louis, C., eds.), Chapman and Hall, London, pp 67–91 13 Sinden, R E., Suhrbier, A., Davies, C S., Fleck, S L., Hodivala, K., and Nicholas, J C (1990)... Sinden, R E and Croll, N A (1975) Cytology and kinetics of microgametogenesis and fertilization of Plasmodium yoelii nigeriensis Parasitol 70, 53–65 22 Janse, C J and Waters, A P (1995) Plasmodium berghei: the application of cultivation and purification techniques to molecular studies of malaria parasites Parasitol Today 11, 138–143 23 Al-Olayan, E., Beetsma, A L., Butcher, G A., Sinden, R E., and Hurd,... continuous function of parasite density (16) and can be further extended to include other covariates (17) Schellenberg et al (18) used the fraction of fever cases in a population attributable to malaria at each level of parasite density to evaluate the sensitivity and specificity of alternative case definitions for malaria, and to provide a direct estimate of malariaattributable fever The attributable... of malaria and define their utility in the context of local parameters of endemicity, infrastructure, and intent of inquiry The risk of malaria is highly dependent on interactions between the host, parasite, mosquito vector, and environment, a relationship known as the epidemiologic triad of disease Changes in any one of these elements may profoundly impact risk of infection Measures of risk of malaria . by Denise L. Doolan Malaria Methods and Protocols Humana Press Humana Press Edited by Denise L. Doolan Malaria Methods and Protocols Vector Incrimination and EIR 3 3 From: Methods in Molecular. dipping methods. Mosquitoes are identified by standard taxo- nomic methods and also by molecular methods if mosquitoes belong to a species complex. The standard methods for performing landing/biting. the glands become stuck in the mosquito thorax, and it is necessary to tease apart the tissue to locate and cut the glands. 5. After severing the salivary glands, remove the head and thorax and