a new pcr assay for simultaneous studies of ...

A NEW PCR ASSAY FOR SIMULTANEOUS STUDIES OF LEUCOCYTOZOON, PLASMODIUM, AND HAEMOPROTEUS FROM AVIAN BLOOD Author(s): Olof Hellgren, Jonas Waldenström, Staffan Bensch Source: Journal of Parasitology, 90(4):797-802. 2004. Published By: American Society of Parasitologists DOI: http://dx.doi.org/10.1645/GE-184R1 URL: http://www.bioone.org/doi/full/10.1645/GE-184R1

BioOne (www.bioone.org) is a nonprofit, online aggregation of core research in the biological, ecological, and environmental sciences. BioOne provides a sustainable online platform for over 170 journals and books published by nonprofit societies, associations, museums, institutions, and presses. Your use of this PDF, the BioOne Web site, and all posted and associated content indicates your acceptance of BioOne’s Terms of Use, available at www.bioone.org/page/terms_of_use. Usage of BioOne content is strictly limited to personal, educational, and non-commercial use. Commercial inquiries or rights and permissions requests should be directed to the individual publisher as copyright holder.

BioOne sees sustainable scholarly publishing as an inherently collaborative enterprise connecting authors, nonprofit publishers, academic institutions, research libraries, and research funders in the common goal of maximizing access to critical research.

J. Parasitol., 90(4), 2004, pp. 797–802 q American Society of Parasitologists 2004

A NEW PCR ASSAY FOR SIMULTANEOUS STUDIES OF LEUCOCYTOZOON, PLASMODIUM, AND HAEMOPROTEUS FROM AVIAN BLOOD Olof Hellgren, Jonas Waldenstro¨m, and Staffan Bensch Department of Animal Ecology, Ecology Building, Lund University, SE-22362 Lund, Sweden. e-mail: [email protected] ABSTRACT: Many bird species host several lineages of apicomplexan blood parasites (Protista spp., Haemosporida spp.), some of which are shared across different host species. To understand such complex systems, it is essential to consider the fact that different lineages, species, and families of parasites can occur in the same population, as well as in the same individual bird, and that these parasites may compete or interact with each other. In this study, we present a new polymerase chain reaction (PCR) protocol that, for the first time, enables simultaneous typing of species from the 3 most common avian blood parasite genera (Haemoproteus, Plasmodium, and Leucocytozoon). By combining the high detection rate of a nested PCR with another PCR step to separate species of Plasmodium and Haemoproteus from Leucocytozoon, this procedure provides an easy, rapid, and accurate method to separate and investigate these parasites within a blood sample. We have applied this method to bird species with known infections of Leucocytozoon spp., Plasmodium spp., and Haemoproteus spp. To obtain a higher number of parasite lineages and to test the repeatability of the method, we also applied it to blood samples from bluethroats (Luscinia svecica), for which we had no prior knowledge regarding the blood parasite infections. Although only a small number of different bird species were investigated (6 passerine species), we found 22 different parasite species lineages (4 Haemoproteus, 8 Plasmodium, and 10 Leucocytozoon).

Species of the apicomplexans Haemoproteus, Plasmodium, and Leucocytozoon comprise a diverse group of vector-transmitted parasites that infect red blood cells (in the case of Leucocytozoon spp., also white blood cells) and other organs within their vertebrate hosts (Atkinson and Van Riper, 1991; Valkiunas, 1993). Species of these parasite genera share several characters with human malaria parasites, and all 3 (but most often only Plasmodium spp.) are referred to as avian malaria. These parasites have served as model organisms for studies on many aspects of parasite–host interactions, including parasite–host evolution (Perkins and Schall, 2002; Ricklefs and Fallon, 2002), host life-history trade-offs (Richner et al., 1995; Nordling et al., 1998), and sexual selection (Hamilton and Zuk, 1982). The frequent use of these parasites in evolutionary and population ecology studies is based on the relative ease with which infected birds can be distinguished from uninfected ones and the fact that the intensity of infection can be estimated for each host using blood smears (Valkiunas, 1993; Richner et al., 1995; Rintama¨ki et al., 1998). Thus, both quantitative and qualitative methods can be applied to examine the costs of infection. Species of Haemoproteus, Plasmodium, and Leucocytozoon are closely related genetically but differ in life-history traits. The main body of published work has focused on species of the more easily detected Haemoproteus and Plasmodium, with relatively few studies on Leucocytozoon spp. (Atkinson and Van Riper, 1991). The scarcity of investigations on Leucocytozoon spp. infections is not because the infection is itself rare but because the life stages of Leucocytozoon spp. are detectable in peripheral blood for only very short time periods, which makes the infection difficult to detect and accurately identify using traditional ocular methods (Fallis and Desser, 1977; Valkiunas, 1997). Despite the methodological difficulties, Leucocytozoon spp. have been found to be a common (sometimes the most common) parasite in some bird populations, primarily in the temperate regions of the Northern Hemisphere (Rintama¨ki et al., 1998; Deviche et al., 2001). Some of the problems associated with the traditional typing of blood parasites can now be solved with molecular methods,

which are generally much more sensitive than traditional microscopic procedures (Perkins et al., 1998; Richard et al., 2002). Furthermore, polymerase chain reaction (PCR)–based methods can provide sequence information that allows for the identification of the specific parasite lineage below the species level ˚ kesson, 2003), which is not possible using par(Bensch and A asite morphology alone. There are several published PCR-based methods (Bensch et al., 2000; Perkins and Schall, 2002; Ricklefs and Fallon, 2002; Fallon et al., 2003) and 1 serological technique (Atkinson et al., 2001) for the detection and identification of Plasmodium spp. and Haemoproteus spp. from birds. However, until now, there has been no general protocol available for the detection of Leucocytozoon spp. The cytochrome b gene of the apicomplexan parasite mitochondrial genome has been found to have conserved regions for construction of primer sites with variable sections of DNA between the conserved regions, which have made it suitable for detection and identification of Haemoproteus and Plasmodium lineages (Waldenstro¨m et al., 2004). In this article, we describe a nested-PCR assay, which targets the cytochrome b gene of the parasites, thus enabling the screening and typing of Leucocytozoon species in parallel with species of Haemoproteus and Plasmodium in avian blood samples. The protocol involves a first PCR step, modified from Waldenstro¨m et al. (2004), which amplifies parasite DNA from all 3 genera, and then a choice of 2 primer pairs to either amplify Leucocytozoon spp. singly or to amplify Haemoproteus spp. and Plasmodium spp. together (Haemoproteus–Plasmodium). This new method appears highly repeatable and reliable and provides an important tool for simultaneous studies of species of Haemoproteus, Plasmodium, and Leucocytozoon in birds. Moreover, this method also provides the possibility of sequence-based identification of different mitochondrial lineages within the parasite genera. MATERIALS AND METHODS Primer design Primers were designed within the 59 end of the cytochrome b gene of the parasite mitochondrial genome by using the published sequences of avian Haemoproteus, Plasmodium, and Leucocytozoon mitochondrial

Received 2 July 2003; revised 24 November 2003; accepted 24 November 2003. 797

798

THE JOURNAL OF PARASITOLOGY, VOL. 90, NO. 4, AUGUST 2004

FIGURE 1. Schematic illustration of the directions and combinations of the different primers. 1. Primers for amplification of the initial step in the nested PCR. 2. Primer combination for amplification of Haemoproteus and Plasmodium lineages. 3. Primer combination for amplification of Leucocytozoon lineages. Note that the primer HaemR2/R2L/ NR3 is the reversed compliment and is shown as it should be ordered (59–39). Underlined letters indicate the primer sequences not shared between HaemF–HaemFL and HaemR2–HaemR2L. I stands for a universal base, inosine.

DNA (mtDNA) (Perkins and Schall, 2002). Primer pairs that amplify the conserved regions of the Haemoproteus spp. and Plasmodium spp. cytochrome b genes have previously been developed at Lund (Bensch et al., 2000; Waldenstro¨m et al., 2004). In the original protocol (Bensch et al., 2000), the HaemF and HaemR2 primers were used to amplify a 480-bp fragment (excluding primers) with a single PCR. The performance of this PCR was significantly improved, especially for the detection of low-intensity Plasmodium sp. infections, by extending the single PCR to a nested PCR (for full details see Waldenstro¨m et al., 2004). HaemNF and HaemNR2 primers were used to initially amplify a 617-bp large fragment (including the primers), to which the HaemF and HaemR2 primers of Bensch et al. (2000) could be internally nested in a second PCR step. Based on sequence homology among aligned sequences of the parasite genera, initial primers were constructed (HaemNFI [59-CATATATTAAGAGAAITATGGAG-39] [I 5 a universal base, inosine] and HaemNR3 [59-ATAGAAAGATAAGAAATACCATTC-39]) to amplify parasite mtDNA from species of Haemoproteus, Plasmodium, and Leucocytozoon. For the second PCR, we used HaemF–HaemR2 primers (Bensch et al., 2000) for Plasmodium spp. and Haemoproteus spp. and constructed 2 new primers, HaemFL (59-ATGGTGTTTTAGATACTT ACATT-39) and HaemR2L (59-CATTATCTGGATGAGATAATGGIG C-39) for Leucocytozoon spp. (Fig. 1). The first PCR was performed in volumes of 25 ml, which included 50 ng of total genomic DNA, 1.25 mM of each deoxynucleoside triphosphate, 1.5 mM MgCl2, 13 PCR (Applied Biosystem, Foster City, California), 0.6 mM of each primer, and 0.5 units Taq DNA polymerase. The PCRs including HaemNFI–HaemNR3 were conducted using the following conditions: 30 sec at 94 C, 30 sec at 50 C, and 45 sec at 72 C for 20 cycles. The samples were incubated before the cyclic reaction at 94 C for 3 min and after the cyclic reaction at 72 C for 10 min. We used 2 ml of the first PCR reaction as the template for the second PCR, 1 ml for Leucocytozoon spp. (HaemFL–HaemR3L) and 1 ml for Haemoproteus spp.–Plasmodium spp. (HaemF–HaemR2). These PCRs were performed separately in 25-ml volumes with the same proportions of reagents as in the initial PCR reactions. The thermal profile of the PCR was identical to the initial PCR but performed for 35 cycles instead of 20 cycles. To check if the PCRs had been amplified successfully, we ran 1.5 ml of the final PCR product on a 2% agarose gel and used samples with positive amplification for further evaluation. Evaluation To evaluate the ability of the method to detect Leucocytozoon spp. infections, we used samples from birds with known infections of different parasites, i.e., where infection had been confirmed morphologically using Geimsa-stained blood smears. The test panel included birds with single infections of Leucocytozoon spp. (2 juvenile bramblings, Fringilla montifringilla) and birds with simultaneous infections of both Haemoproteus spp. and Leucocytozoon spp. (1 bluetit, Parus caereu-

leus; 1 crossbill, Loxia curvirostra; 1 robin, Erithacus rubicula; and 1 siskin, Carduelis spinus). To further test the new method, we applied the nested-PCR assays on a collection of blood samples (n 5 86) from adult bluethroats for which we had no prior parasite infection knowledge. Each blood sample contained 5–20 ml of blood and had been stored in SET buffer (0.015 M NaCl, 0.05 M Tris, 0.001 M ethylenediaminetetraacetic acid, pH 8.0) at ambient temperature in the field and later at 220 C. DNA was extracted using a standard chloroform–isoamylalcohol method (Sambrook et al., 2002), and diluted genomic DNA was used as template in the PCR assays. Samples showing positive amplification were selected for sequencing using procedures as described by Bensch et al. (2000). Fragments were first sequenced from the 59 end with either HaemF (in the case of Haemoproteus-Plasmodium spp.–positive samples) or HaemFL (in the case of Leucocytozoon spp.–positive samples), and the obtained sequences were edited and aligned using the program BioEdit (Hall, 1999). Samples with positive amplification yielded PCR products of 478 bp (excluding primers) for Leucocytozoon spp. and 480 bp for Haemoproteus spp. and Plasmodium spp. All unique haplotypes, i.e., sequences differing by 1 or more base pair from any of the other obtained sequences, were sequenced from the 39 end using HaemR2 (Haemoproteus-Plasmodium spp.–positive samples) or HaemR2L (Leucocytozoon spp.–positive samples) for sequence validation. In cases where mixed infections occurred (n 5 3), observed as ‘‘double base calling’’ in the electropherogram, fragments were cloned and separated using a TA-cloning kit (Invitrogen, Carlsbad, California) according to the manufacturer’s instructions. We amplified the inserted DNA from 10 colonies per plate using standard M13 primers and sequenced from 1 direction using the forward primer. We used the program MEGA, version 2.1 (Kumar et al., 2001), and the neighbor-joining method with a Kimura 2-parameter distance matrix to investigate the genetic relationship of the parasite sequences obtained in this study with published sequences. The tree was rooted with Theileria annulata (Piroplasmidia). Repeatability and detection The repeatability of the method was estimated by running 30 bluethroat DNA samples 3 times for both the Leucocytozoon and the Plasmodium–Haemoproteus samples. The outcomes of the different runs were compared using a 1-way analysis of variance to calculate amongand within-individual variation of scored prevalence (Lessells and Boag, 1987). To further evaluate the success of the amplification of Haemoproteus–Plasmodium spp. and to determine if the performance differed between the 2 nested-PCR methods, the same samples were tested using primers developed for the exclusive amplification of Plasmodium spp. and Haemoproteus spp. (Waldenstro¨m et al., 2004). The detection success of the method was tested with a dilution series as in Fallon et al. (2003). Three dilution series stemming from each of 2 birds with single infections of Haemoproteus and Plasmodium and 1 with Leucocytozoon spp. were tested for positive amplification. The samples were diluted with uninfected bird DNA, keeping the total concentration at 25 ng/ml, in steps of 10, down to a 1,000,000-fold dilution (Table I). The original intensity of the infection was scored under a light microscope as in Fallon et al. (2003).

RESULTS Detection of apicomplexan parasite infections The nested PCR for the detection of Leucocytozoon spp. correctly amplified all samples in the test panel where Leucocytozoon or Leucocytozoon and Haemoproteus parasites had been recorded by microscopy. Among the test panel of birds with unknown infections, the PCR assays gave an overall prevalence of apicomplexan parasites in 59% (n 5 86) of birds, with varying genus-specific prevalences, i.e., 24% of 8 lineages for Plasmodium, 48% of 5 lineages for Leucocytozoon, and 1.2% of a single lineage for Haemoproteus (Fig. 2). The method also identified several individuals with simultaneous infections with

HELLGREN ET AL.—SIMULTANEOUS STUDY OF 3 PARASITE GENERA

799

TABLE I. Dilution series of malaria-infected birds. Each infected bird was tested for positive PCR amplification in three dilution series. Limits of infection intensities for consistent scoring (all positive) and minimum detection (at least one positive) are calculated based on smear intensities. Prefix in the lineage name corresponds to h 5 Haemoproteus, p 5 Plasmodium, and l 5 Leucocytozoon.

Lineage

Smear intensity (%)

Number of positive amplifications (max 3) 1

1021

1022

1023

1024

1025

hGRW1 hGRW1 pGRW2 pGRW2 1Bram1

3.75 2.4 1.16 0.022 0.03

3 3 3 3 3

3 3 3 3 3

3 3 3 2 2

3 3 3 0 1

1 0 2 0 0

0 0 0 0 0

species from 2 parasite genera (10% with Leucocytozoon–Plasmodium infections and 1.2% with Leucocytozoon–Haemoproteus) or with 2 lineages from the same parasite genus (2 individuals with 2 lineages of Leucocytozoon and 1 individual with 2 lineages of Plasmodium). The repeatability of both methods was high. Of the 30 samples tested with the Plasmodium and Haemoproteus primers, 17

1026

Consistent scoring limit (%)

Minimum detection limit (%)

0 0 0 0 0

0.00375 0.0024 0.00116 0.0022 0.003

0.00038 0.00024 0.00012 0.00022 0.00003

were negative and 9 positive in all 3 runs, whereas 4 samples showed mixed positive and negative results (R 5 0.814, F1,89 5 14.147, P , 0.0001; Fig. 3). The Leucocytozoon spp. primers gave identical results in all 3 runs for 25 of the 30 tested samples (14 consistently negative and 11 positive), whereas 5 samples showed mixed positive and negative results (R 5 0.740, F1,89 5 9.546, P , 0.0001; Fig. 3). The amplification using the

FIGURE 2. Genetic relationship (neighbor joining and a Kimura 2-parameter distance method) of blood parasites. Bootstrap values presented on the clade branches. Shaded lineages are obtained from previous studies or from GenBank. Underlined lineages were found in a sample set of bluethroats (Luscinia svecica) with unknown infections. † indicate lineages found in control birds with known infections. The tree was rooted with Theileria annulata (Piroplasmidia).

800


DISCUSSION Detection of parasite lineages

FIGURE 3. Number of positive or negative amplification of blood parasites in 3 runs of 30 test individuals. Each individual bird was tested 3 times. Black, individuals tested for Leucocytozoon spp.; white, individuals tested for Plasmodium spp.–Haemoproteus spp.

primers from Waldenstro¨m et al. (2004) did not yield any new positive samples among the 30 tested. Our nested-PCR method provided positive amplifications in all cases down to dilutions corresponding to 1 parasite per 100,000 host blood cells for Haemoproteus, Plasmodium, and Leucocytozoon (Table I). In dilutions corresponding to 1 parasite per 1,000,000 host blood cells, 50% of the samples in the Haemoproteus spp. dilution series, 67% in the Plasmodium spp. series, and 67% in the Leucocytozoon spp. series showed positive amplification. The method did not show any positive amplification for less than 1 parasite per 1,000,000 host blood cells except 1 positive case for Leucocytozoon at the dilutions corresponding to 1 parasite per 10,000,000 host blood cells (Table I). Phylogenetic relationships The sequenced PCR products of the 2 PCR assays were edited and aligned, and the resulting alignment of 480 bp was analyzed. The sequences showed high variability, and a neighbor-joining tree clustered them into 3 distinct groups with high confidence (bootstrap values from 88 to 98 for the different genera and branches; Fig. 2). These 3 clusters corresponded to species of Haemoproteus, Plasmodium, and Leucocytozoon. Each cluster was confirmed using published genus-identified sequences, together with the tested samples of known infections from this study. All sequences obtained with the Leucocytozoon spp. primer combination grouped in the Leucocytozoon cluster, and all the sequences obtained with the Haemoproteus spp. and Plasmodium spp. primer pair grouped with known Haemoproteus and Plasmodium sequences. Furthermore, apart from providing generic identification of tested samples, the obtained fragment of the cytochrome b gene was also phylogenetically informative within genera, as observed in large lineage variation in all 3 obtained clusters. Among the sequences included in this analysis, the deepest split was 6.3% within Haemoproteus spp., 19.2% within Plasmodium spp., and 22.8% within Leucocytozoon spp.

Several PCR-based methods for studies of Haemoproteus spp. and Plasmodium spp. have been published recently (Bensch et al., 2000; Ricklefs and Fallon, 2002; Fallon et al., 2003), but the present investigation reports the first protocol for a general detection of Leucocytozoon spp. PCR-based methods are more sensitive than traditional microscopy (Richard et al., 2002); however, performance differs markedly between methods. In a comparative study, Richard et al. (2002) concluded that PCR assays targeting parasite mtDNA had superior performance over those targeting nuclear ribosomal genes. In our laboratory, we have developed a nested-PCR assay (Waldenstro¨m et al., 2004), based on the primer pairs published in Bensch et al. (2000), which further improved the detection capability, especially for low-intensity Plasmodium spp. infections. In this study, we have further developed the methods of Bensch et al. (2000) and Waldenstro¨m et al. (2004) by making initial primers that include Leucocytozoon spp. mtDNA in the amplification and an additional set of primers for the second step that allow for separation of Leucocytozoon spp. infections. Thus, from extracted blood samples, only 3 PCR runs are needed to identify individuals infected with species of Plasmodium– Haemoproteus or Leucocytozoon in a study population of birds. The new method correctly identified all samples with known infections, and the repeatability was high for the detection of Plasmodium spp., Haemoproteus spp., and Leucocytozoon spp. Because the methods are performed on DNA extracted from blood samples, the prerequisite for successful amplification is the presence of parasites in the circulating blood of the host. Because results are obtained as the presence or absence of bands in an agarose gel, a validation of the total DNA quality, e.g., by first using molecular sexing of the hosts, is needed before trusting negative scores in the malaria PCR. In the case of the bluethroat with unknown infections, the blood samples had been previously used for host microsatellite and amplified fragment length polymorphism typing (data not shown). Although the repeatability of parasite detection was high, some variation was observed between trials. The obtained variation was most likely associated with low parasitemia because variation also occurred at very low intensities when the method was tested for intensity dependence. However, the method allows detection of infections without failures in the dilutions corresponding to 1 parasite per 100,000 host blood cells, and inconsistent results were obtained initially in dilutions corresponding to 1 parasite per 1,000,000 host blood cells (Table I). Hence, the method successfully identified infections with intensities of $0.001% infected blood cells, with the possibility of identifying infections with intensities of $0.0001% infected blood cells. The occurrence of the variation at very low intensities is likely to be due to chance events during the first PCR cycles that determine whether a fragment will be amplified or not. Even though the repeatability is high and the detection rate is roughly twice that of ocular investigation (Waldenstro¨m et al., 2004), studies of processes (fitness) at the individual level should consider using repeated screening to include and increase the reliability of birds with the lowest infection intensities.

HELLGREN ET AL.—SIMULTANEOUS STUDY OF 3 PARASITE GENERA

Phylogenetic analysis The apicomplexan cytochrome b gene is phylogenetically informative and has been used previously to establish the evolution and divergence of avian malarial parasites in different host families (Perkins and Schall, 2002; Ricklefs and Fallon, 2002) and to establish the areas in which these parasites are transmitted (Waldenstro¨m et al., 2002). This study clearly shows that the cytochrome b gene is also suitable for constructing well-supported phylogenies for Leucocytozoon species. Our method provided highly variable sequences with the largest observed sequence divergence of 23% between Leucocytozoon dubreuli (AY099064) and Leucocytozoon sp. (AY465560) (Fig. 2). The largest divergence between lineages found in the same host species was 8.4% (between lBT1 and lBT2). This is a deeper divergence than that between Plasmodium falciparum (found in humans) and P. reichenowi (found in chimpanzee) (3.3%), which are supposed to have diverged approximately 5 million yr ago (Escalante et al., 1998). Thus, the most divergent lineages of Leucocytozoon spp. encountered in the same host species most likely reflect different parasite species. On the other hand, 2 of the obtained Leucocytozoon lineages, lBT5 and lBT4, differed by only 1 and 2 substitutions, respectively, from 2 other lineages (lBT1 and lBT2, respectively), possibly indicating intraspecies mtDNA variation (Fig. 2). The construction of a powerful method for the detection and identification of Leucocytozoon spp., in parallel with the method for Plasmodium spp. and Haemoproteus spp., will hopefully open the way for studies on the occurrence and effect of these parasites in natural populations of birds. To neglect the contribution of fitness effects stemming from Leucocytozoon spp. infections could confound results, especially in northern temperate areas where Leucocytozoon spp. and its main vector, the blackfly (Simuliidae), are both common (Malmqvist, 1994; Malmqvist et al., 1999; Deviche et al., 2001; Ojanen et al., 2002). The relative importance of Leucocytozoon spp. infections is illustrated by the bluethroats screened in this study, i.e., Leucocytozoon spp. infections were twice as common as Plasmodium spp. (48% and 24% prevalence, respectively). Therefore, a study focusing only on Haemoproteus spp. and Plasmodium spp. parasites might give a biased result because it would leave out the most common blood parasite species. Moreover, combinations of parasites may have different effects on host fitness because they may, positively or negatively, interact with each other (Riche, 1988). ACKNOWLEDGMENTS The study was supported by grants from Stiftelsen Lunds Djurskyddsfond and Olle och Signild Engkvist Stiftelser. Thanks to Charlotte Val˚ ke Lindstro¨m for makind for excellent help during the sampling trip, A ing the field trip possible, Kjell Grimsby, Thomas Holmberg, Trond ˚ ke Andersson for help during the field season. Amundsen, and Nils-A We thank Dennis Hasselquist for scientific input, Gediminas Valkiunas for providing blood samples, and Martin Stjernman for evaluating infection intensities. David Richardson, Gediminas Valkiunas, and Ravinder Seghal provided insightful comments on the manuscript.

LITERATURE CITED ATKINSON, C. T., R. J. DUSEK, AND J. K. LEASE. 2001. Serological responses and immunity to superinfection with avian malaria in experimentally-infected Hawaii Amakihi. Journal of Wildlife Diseases 37: 20–27.

801

———, AND C. VAN RIPER III. 1991. Pathogenicity and epizootiology of avian haematozoa: Plasmodium, Leucocytozoon, and Haemoproteus. In Bird-parasite interaction. J. T. Loye and M. Zuk (eds.). Oxford University Press, Oxford, U.K., p. 19–48. ˚ KESSON. 2003. Temporal and spatial variation of BENSCH, S., AND S. A hematozoans in Scandinavian willow warbler. Journal of Parasitology 89: 388–391. ¨. O ¨ STMAN, B. HANSSON, H. ———, M. STJERNMAN, D. HASSELQUIST, O WESTERDAHL, AND R. T. PINHEIRO. 2000. Host specificity in avian blood parasites: A study of Plasmodium and Haemoproteus mitochondrial DNA amplified from birds. Proceedings of the Royal Society of London Series B—Biological Sciences 267: 1583–1589. DEVICHE, P., E. C. GREINER, AND X. MANTECA. 2001. Seasonal and agerelated changes in blood parasite prevalence in dark-eyed Juncos (Junco hyemalis, Aves, Passeriformes). Journal of Experimental Zoology 289: 456–466. ESCALANTE, A. A., D. E. FREELAND, W. E. COLLINS, AND A. A. LAL. 1998. The evolution of primate malaria parasites based on the gene encoding cytochrome b from the linear mitochondrial genome. Proceedings of the National Academy of Sciences of the United States of America 95: 8124–8129. FALLIS, M. A., AND S. S. DESSER. 1977. On species of Leucocytozoon, Haemoproteus and Hepatocystis. In Parasitic protozoa, J. P. Kreier (ed.). Academic Press, New York, p. 239–266. FALLON, S. M., R. E. RICKLEFS, B. L. SWANSON, AND E. BERMINGHAM. 2003. Detecting avian malaria: An improved polymerase chain reaction diagnostic. Journal of Parasitology 89: 1044–1047. HALL, T. A. 1999. BioEdit: A user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acid Symposium Series 41: 95–98. HAMILTON, W. D., AND M. ZUK. 1982. Heritable true fitness and bright birds—A role for parasites. Science 218: 384–387. KUMAR, S., T. KOICHIRO, I. B. JACOBSEN, AND M. NEI. 2001. MEGA2: Molecular evolutionary genetics analysis software. Bioinformatics 17: 1244–1245. LESSELLS, C. M., AND P. T. BOAG. 1987. Unrepeatable repeatabilities— A common mistake. Auk 104: 116–121. MALMQVIST, B. 1994. Preimaginal blackflies (Diptera, Simuliidae) and their predators in a central Scandinavian lake outlet stream. Annales Zoologici Fennici 31: 245–255. ———, Y. X. ZHANG, AND P. H. ADLER. 1999. Diversity, distribution and larval habitats of North Swedish blackflies (Diptera: Simuliidae). Freshwater Biology 42: 301–314. NORDLING, D., M. ANDERSSON, S. ZOHARI, AND L. GUSTSFSSON. 1998. Reproductive effort reduces specific immune response and parasite resistance. Proceedings of the Royal Society of London Series B— Biological Sciences 265: 1291–1298. OJANEN, U., O. RATTI, P. H. ADLER, K. KUUSELA, B. MALMQVIST, AND P. HELLE. 2002. Blood feeding by black flies (Diptera: Simuliidae) on black grouse (Tetrao tetrix) in Finland. Entomologica Fennica 13: 153–158. PERKINS, S. L., S. M. OSGOOD, AND J. J. SCHALL. 1998. Use of PCR for detection of subpatent infections of lizard malaria: Implications for epizootiology. Molecular Ecology 7: 1587–1590. ———, AND J. J. SCHALL. 2002. A molecular phylogeny of malarial parasites recovered from cytochrome b gene sequences. Journal of Parasitology 88: 972–978. RICHARD, F. A., R. N. M. SEHGAL, H. I. JONES, AND T. B. SMITH. 2002. A comparative analysis of PCR-based detection methods for avian malaria. Journal of Parasitology 88: 819–822. RICHE, T. L. 1988. Interaction between malaria parasites infecting the same vertebrate host. Parasitology 96: 607–639. RICHNER, H., P. CHRISTE, AND A. OPPLIGER. 1995. Paternal investment affects prevalence of malaria. Proceedings of the National Academy of Sciences of the United States of America 92: 1192–1194. RICKLEFS, R. E., AND S. M. FALLON. 2002. Diversification and host switching in avian malaria parasites. Proceedings of the Royal Society of London Series B—Biological Sciences 269: 885–892. RINTAMA¨KI, P. T., W. OJANEN, H. PAKKALA, AND M. TYNJA¨LA¨. 1998. Blood parasites of migrating willow warblers (Phylloscopus trochilus) at a stopover site. Canadian Journal of Zoology—Revue Canadienne de Zoologie 76: 984–988. SAMBROOK, J., F. J. FRITCH, AND T. MANIATIS. 2002. Molecular cloning,

802


a laboratory manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. VALKIUNAS, G. 1993. Phatogenic influence of haemosporidians and trypanosomes on wild birds in the field conditions: Facts and hypothesis. Ekologija 1: 47–60. ———. 1997. Bird Haemosporida. Institute of Ecology, Vilnius, Lithuania, 608 p.

¨. O ¨ STMAN. 2004. WALDENSTRO¨M, J., S. BENSCH, D. HASSELQUIST, AND O A new nested polymerase chain reaction method very efficient in detecting Plasmodium and Haemoproteus infections from avian blood. Journal of Parasitology 90: 191–194. ———, ———, S. KIBOI, D. HASSELQUIST, AND U. OTTOSSON. 2002. Cross-species infection of blood parasites between resident and migratory songbirds in Africa. Molecular Ecology 11: 1545–1554.

J. Parasitol., 90(4), 2004, p. 802 q American Society of Parasitologists 2004

BOOK REVIEW . . . Parasites and Diseases of Wild Birds in Florida, by D. J. Forrester and M. G. Spalding. University Press of Florida, Gainesville, Florida. 2003. 1132 p. Cloth cover, ISBN: 0-8130-2560-5. At first glance, a book with the above title might not seem to be of great general interest. However, Florida has a highly diverse bird fauna, the state is a major migration site, and the authors have worked with the Florida Fish and Wildlife Conservation Commission for decades, assessing the effect of parasites and disease on both birds and mammals. As a result, this book is one of the most comprehensive, easily used, and data-rich resources available to parasitologists. It will be especially relevant, indeed virtually a required reference volume, for those in any area of conservation biology or wildlife management, and it will be very useful to teachers and researchers who are working with natural host–parasite systems involving birds. Parasites and Diseases of Wild Birds in Florida is organized more or less along classical taxonomic lines, with chapters on bird families beginning with loons and grebes and ending with passerines. ‘‘Parasite’’ and ‘‘disease’’ are both interpreted broadly, thus information is provided not only on viruses, bacterial infections, fungi, protozoa, helminths, and arthropods but also on chemical residues, e.g., lead poisoning and organophosphates, and trauma, e.g., injury resulting from contact with structures. The book is heavily referenced (even the Preface has a literature cited section!) and as a result is a virtual window revealing a vast body of literature, not typically accessible to the average academic or government employee. For example, the 22-page chapter on kites has nearly 4 pages of references, with citations ranging from those in major journals to ones in relatively obscure commission reports. Each chapter also is generously supplied with tables, including data sources,

an excellent example being a 4-page table on ticks reported from passerines, with collection sites, dates, tick stages represented, and prevalence. There are 23 tables listing similar data for helminth parasites of wild turkeys. The tabular data and Literature Cited alone make this single volume an exceptionally valuable resource, potentially saving enormous amounts of time for anyone who has any reason whatsoever to recover information on bird diseases or parasites. This book is also unusual in that it mentions both negative data and gaps in our knowledge, although the information applies mainly to Florida. For example, in the chapter on Anserinae (whistling ducks, swans, and geese), we are told there is no information on neoplasia, viruses, bacteria, fungi, or blood protozoa in these (wild) birds, but we are also told of reports on some of those parasites in captive flocks or in nearby states. Ecological data are included when available and relevant, a good example being seasonal dynamics of nematodes in bobwhites. The book provides a reasonable number of photographs of pathological conditions, some of which are quite dramatic, thus useful in teaching. Techniques are also illustrated, and although some readers might question why these photographs are included, it is not always obvious to biologists in general, especially in a molecular age, how data on wild animals are acquired. A good example of this material is the series of photographs on how to make and use a lard-can bait trap to get information on avian malaria vectors. The text is well written. The David Maehr pen and ink bird drawings are a nice touch for a scientific book, and the originals of these drawings are probably collectibles because of their subtle quality. John Janovy, Jr., School of Biological Sciences, 348 Manter Hall, University of Nebraska–Lincoln, Lincoln, Nebraska 68588-0118.