America’s Environmental Report Card: Are We Making the Grade?

By Harvey Blatt 
 
Cambridge, MA:MIT Press, 2005. 277 pp. ISBN: 0-262-02572-8, $27.95 cloth 
 
Protecting the environment continues to be a top priority for most Americans. Some examples of major environmental concerns include water and air pollution, hazardous waste disposal, use of toxic chemicals, and environmental threats to children. In America’s Environmental Report Card: Are We Making the Grade? Harvey Blatt provides a comprehensive status report on the following nine selected complex environmental problems: water pollution, dangers of floods, leaching of garbage from landfills, pesticide runoff, depletion of energy resources, global warming, air pollution, ozone depletion, and radiation from nuclear power plants and the storage of nuclear wastes. He also describes whether the situation with respect to these issues is deteriorating or improving and what actions can be taken at the individual, corporate, and political levels to ensure safe and adequate resources for future generations. 
 
This book contains a large amount of information and statistics, often presented in charts, figures, and tables. It is easily comprehensible, and one does not “get lost in the numbers.” Its conversational tone, interspersed with anecdotes and humor, makes the 238 pages of text easy to read and at times entertaining. 
 
However, the book does not provide a scientific review or analysis of the selected environmental issues and at times does not provide a balanced scientific viewpoint. For example, the introductory statement that “data suggesting that toxic agents in the environment have reduced the average male sperm count by 42 percent in the past 50 years” is not backed up by references nor is there mention of other references that do not support this hypothesis. Such a statement in isolation could be very alarming to the general public. The references cited in the book are mainly secondary sources, in many instances from magazine and newspaper articles. Perhaps it is this reliance on secondary sources that sometimes leads to inaccuracies and misleading statements. For example, there is no scientific evidence to indicate that global warming may result in an “epidemic” of an increase in male babies over female (p. 148). Similarly, respiratory disease is not the greatest killer of children on the planet (p. 155); diarrheal and infectious diseases are the major contributors to global child mortality. 
 
The author, perhaps unintentionally, provides somewhat of a doomsday scenario for a number of the environmental problems covered. For example, at the beginning of the chapter of air pollution (Chapter 7) the reader is inundated with detailed descriptions and disturbing statistics on the morbidity and mortality resulting from exposure to air pollutants. Only at the end of the chapter is there a short paragraph noting that “air quality has improved markedly … since the passage of the Clean Air Act in 1972” (p. 175). Similar observations can be made for several of the other chapters. 
 
Unfortunately, the title of this book is misleading. On the book jacket, actual grades (e.g., B, C) are given to the nine issues covered in the text. I had anticipated finding out how these grades were derived—what criteria and standards were used. Nowhere in the text is there any discussion of a “report card” or how the grades on the book jacket were determined. A concluding paragraph for each chapter provides a limited subjective evaluation on the current status of the environmental issue discussed but does not address whether America is making the grade in any systematic, objective manner. 
 
Nevertheless, those wishing to learn about the very real environmental problems facing the United States will find Blatt’s book very interesting, full of factual information, and eminently readable. It should serve as a valuable resource for the public, government officials, and scientists new to the field. It would also be excellent background reading for a graduate course in environmental sciences.


Background
Plasmodium vivax is the most geographically widespread of the four Plasmodium species infective to humans found throughout South and Central America, Asia, the Middle East, and parts of Africa and infects an estimated 70-80 million people annually [1]. Chloroquine (CQ)-resistant Plasmodium falciparum, and to a lesser extent CQ resistant P. vivax, is almost as endemic as malaria itself and alternatives such as the drug combination sulphadoxine/ pyrimethamine (SP) have replaced CQ. Resistance to SP has recently emerged for P. falciparum, while for P. vivax it has been observed sporadically [2]. The molecular mechanisms involved in the development of SP resistance of the two species are most likely similar [3,4]. In P. falciparum, single nucleotide polymorphisms (SNPs) in codon (c) 51, c59 and c108 of the Pfdhfr gene and in c437 and c540 of Pfdhps gene provide pyrimethamine and sulphadoxine resistance, respectively and these SNPs combined result in high risk of SP treatment failure in vivo [5]. For P. vivax, the picture is more complex because pyrimethamine resistance possibly involve several SNPs [6]. However, some evidence support that resistance is mainly associated with mutations at c58 (S58R, occurring as two SNPs, either AGA (R 1 ) or AGG (R 2 )) and c117 (S117N or S117T) with additional mutations at c57 (F57L-existing as three SNPs, CTC (L 1 ), TTG (L 2 ) and TTA (L 3 )) and c61 (T61M) in the Pvdhfr gene [3,4,[6][7][8]. The quadruple mutant haplotype (57L+58R+61M+117T) has been shown to correlate with SP treatment failure in vivo [8] and increases P. vivax resistance to pyrimethamine by more than 500 times [4,6].
Presumably, P. vivax sulphadoxine resistance is caused by SNPs in the Pvdhps gene. Based on homology models of both P. falciparum and P. vivax DHPS enzymes, Korsinczky et al. predicted that the P. vivax wildtype at c585 (V585) possibly cause some level of innate sulphadoxine resistance, while SNPs at c383 (A383G) and c553 (A553G) in Pvdhps most likely increase resistance levels [9]. Imwong et al. showed that only in regions with high SP usage, SNPs in both Pvdhfr and Pfdhps were observed and, furthermore, parasites harbouring multiple mutations in Pvdhfr and Pvdhps were cleared more slowly from the blood of patients following SP treatment [10]. Therefore, P. vivax SP resistance is most likely measurable by examining the frequency of SNPs in both the Pvdhfr and Pvdhps genes.
In Sri Lanka, CQ plus primaquine (PQ) and SP plus PQ are used as 1 st -and 2 nd -line treatment, respectively, against uncomplicated malaria infections, although CQ resistant P. falciparum infections have been reported since 1984 and P. falciparum SP resistance has been observed recently [11]. P. vivax resistance to either CQ or SP has not been recorded on the island. This study investigated the frequency of SNPs/haplotypes in the Pvdhfr (at c57, 58, 61 and 117) and Pvdhps (at c383, 553 and 585) genes in samples collected from nine districts with endemic P. vivax malaria in Sri Lanka over a 1 1/2-year period. The detection of SNPs/haplotypes in Pvdhfr and Pvdhps was performed by applying a new simple enzyme-linked immunosorbent assay (ELISA) using sequence specific oligonucleotide probes (SSOPs) similar to the method detecting SNPs/haplotypes in P. falciparum dhfr, dhps and crt [12].

Materials and methods
The samples originated from individuals seeking treatment for malaria at government health facilities located in nine different malarious district across Sri Lanka. In Sri Lanka the great majority of individuals with perceived malaria seek treatment at government facilities [13]. Samples were collected by routine staff at the facilities trained by the Anti-Malaria Campaign (AMC) of Sri Lanka from September 2004 to March 2006, thereby including the traditionally malaria peak transmission seasons in January and one lower peak season around July. Finger prick blood from patients with single P. vivax or mixed P. vivax/ P. falciparum infections, diagnosed by microscopy were spotted on filter paper and sealed in individual zip-lock bags. DNA extraction was carried out by the chelex-100 method as described in [12].
As positive controls of the various Pvdhfr SNPs/haplotypes, 8 P. vivax dhfr allele samples, kindly provided by Carol Sibley (Dept. of Genome Sciences, University of Washington) were used [4]. These represent each of the ten most common c57, 58, 61 and 117 Pvdhfr SNPs/haplotypes. Furthermore, one positive control consisting of a 50 bp DNA fragment was designed mimicking a specific mutated Pvdhfr sequence, comprising the L 3 -mutation in c57 (TTA) and R 2 -mutation in c58 (AGG) (PcL 3 R 2 T in table 1) biotinylated at the 5'-end by the supplier (MWG Biotech, Riskov, Denmark). Likewise for positive controls of Pvdhps 338G and 553G, 50 bp DNA fragments were designed mimicking these specific SNPs (Pc383G and Pc553G in Table 1).
The outer and nested Pvdhfr PCR protocols used are described in [14], with the exception that the reverse nested primer, KH-3R was biotinylated at the 5'-end by the supplier (MWG Biotech, Riskov, Denmark). The outer Pvdhps PCR primers used (PvDHPS-D and PvDHPS-B) and protocols are described by [9]. The nested Pvdhps primers were designed; NL-1 (5'-GCGAGCGTGATTGA-CATC-3') and NR-1-(5'-GCTCATCAGTCTGCACTCC-3') where the reverse primer, NR-1, was biotinylated at the 5'end. The outer and nested Pvdhps PCR were performed as follows: denaturation at 94°C for 2 min followed by 40 cycles of 94°C for 30 sec, 50°C for 30 sec and 65°C for 1 1/2 min and subsequently a 5 min extension step at 65°C. A SSOP-ELISA, similar to the method for SNP/haplotype analysis of P. falciparum dhfr/dhps was developed [12], however using pre-coated streptavidin plates (Nunc, Roskilde, Denmark). The 3'-end digoxigenin-conjugated SSOPs designed to target the most common Pvdhfr and Pvdhps SNP/haplotypes including the time and temperatures in the two rounds of high stringency washing with tetra-methyl-ammonium chloride (TMAC) is given in table 1. Scoring of ELISA data were performed as described elsewhere [12]. To verify the results, Restriction Fragment Length Polymorphism (RFLP) was performed on a subset of the samples using enzymes and methods described by [15]. Polymorphisms in c383 of the Pvdhps gene were identified by digestion with the restriction enzyme HaeIII (New England Biolabs, Medinova, Glostrup, Denmark).
Sequencing was performed on a subset of samples to clarify some of the Pvdhfr and Pvdhps haplotypes; PCR products with A-overhang were cloned into the TOPO TA vector according to manufacturing procedures (Invitrogen), and plasmids were prepared using MiniPrep spin columns (Omega Biotech). Sequencing was done on an ABI Prism 377 (Perkin-Elmer) using the Big Dye terminator reaction mix (Perkin-Elmer).
Ethical clearance for this project was granted by the Committee on Research and Ethical Review at the Faculty of Medicine, Peradeniya, Kandy and verbal consent was obtained from participants, parents and/or guardians.

Results
In the study period, AMC recorded a total of 2717 P. vivax cases in the country, out of which, 2,149 cases came from the nine districts included in this study (79.1%). 454 (21.1%) blood samples from these districts, representing a large range of catchments efficiencies from 7.7% (Monaragala) to 67.5% (Polonnaruwa) were examined. The samples were analysed for SNPs/haplotypes at position c57, 58, 61 and 117 of the Pvdhfr gene and c383, 553 and 585 (only detection of the wildtype V585) of the Pvdhps gene using an array of SSOPs. Samples either repeatedly PCR negative or negative in one or more of the Pvdhfr or Pvdhps codons were omitted from the analysis.
For Pvdhfr, haplotypes could be constructed for 373 samples (84.6 %) including 25 samples with mixed haplotype infections, but where a major haplotype could be deduced (Figure 1). The Pvdhfr FSTS wild haplotype was represented in 257 samples (68.9 %) while the remaining sam-  There was a general tendency for an increase in the frequency of mutant haplotypes late in the study period. However, due to a skewed temporal collection of samples from the various districts the differences was not analysed further.
A subset of samples analysed by sequencing (mainly to confirm the c57L 3 ) and by digestion of c58 and 117 in the Pvdhfr gene by RFLP confirmed the data obtained be the Pvdhfr SSOP-ELISA.
The Pvdhps haplotypes could be constructed for 368 of the 373 Pvdhfr positive samples (98.7 %). Wildtype haplotypes at c383, 553 and 585 (AAV) was seen in 366 of these samples, while two samples from Trincomalee were of the single mutated GAV haplotype. These were confirmed by sequencing. Both samples expressed the double mutated FRTN haplotype in Pvdhfr.

Discussion
The present descriptive study analysed sulphadoxine/ pyrimethamine (SP) resistance-related SNPs in the P. vivax dhfr and Pvdhps genes in samples originating from nine districts in Sri Lanka, a country were both CQ and SP (in combination with primaquine) is still regarded as efficient treatment against uncomplicated P. falciparum and/ or P. vivax infections. The analysis identified six different haplotypes of Pvdhfr while for Pvdhps, only wildtypes were identified except for two cases.
The double mutant haplotype LRTS (F57L, S58R, T61,117S) was the most frequent mutant haplotype and not as expected as a combination of S117N and S58R (FRTN) as observed previously [6,8,16] and in a recent study from India [17]. The P. vivax triple mutant haplotype LRTN, previously found in Thailand and shown to be associated with reduced ability of patients to decrease par-asites ratios [15] was only found once and the quadruple LRMT mutant haplotype causing a high risk of SP treatment failures [8] was not detected, thus indicating that SP (with primaquine) is still efficient against P. vivax infections in Sri Lanka. Nevertheless, it is surprising that almost one third of the tested P. vivax infections were mutated in the Pvdhfr gene, despite that, officially, SP is only used as second-line drug against CQ treatment failures of P. falciparum. A recent study investigating the availability of SP in privately-owned drug vendor shops in Sri Lanka found that SP was virtually absent from the shops [18], thus the specific drug pressure is unlikely to be caused by unauthorized use. More plausible, the mutations are not only an indication of emerging pyrimethamine resistance, but instead reflect the overall antifolate pressure in Sri Lanka. Presently, antifolates such as dapsone, co-trimoxazole and trimethoprim are for instance used against urinary tract infections and chronic bronchitis on the island. Alternatively, similar to development of P. falciparum resistance to pyrimethamine in vivo, P. vivax populations are occasionally exposed to sub-therapeutic levels of pyrimethamine when re-infecting recently SP-treated patients thereby providing optimal conditions for the emergence of SP tolerant P. vivax parasites [19]. Thus, even low drug pressure may facilitate the emergence of drug tolerant/resistant parasites and this may particularly be the case for P. vivax that to a larger extend than P. falciparum possibly can persist in the host unnoticed.
The frequency of Pvdhfr mutant haplotypes was significantly higher in the most Northern regions (Mannar, Vavuniya and Trincomalee) than the rest of the districts examined. This might be indirectly caused by the civil unrest resulting in a shortage of trained medical personnel, non-accurate malaria diagnosis and an underestimation of malaria infections mainly in the Northern part of Sri Lanka [20]. Furthermore, the FRTN haplotype was only observed in the Northern districts and it may be speculated that human migration between Southern India and the Northern part of Sri Lanka has introduced this particular haplotype from the Southern district of Chennai where it is highly prevalent [17]. The limited number of samples received from some districts, e.g. Monaragala, Ampara, Batticaloa, Mannar and Vavuniya, limits interpretation.
The high frequency of mutant haplotypes related to pyrimethamine resistance is worrying because it indicates that drug tolerant/resistant P. vivax parasites have evolved despite a low level of SP drug pressure, possibly attributed to the use of other antifolate drugs. It is not known whether these mutant P. vivax haplotypes do exhibit SP resistance in vivo.