Modulation of Malaria Phenotypes by Pyruvate Kinase (PKLR) Variants in a Thai Population

Rebekah van Bruggen; Christian Gualtieri; Alexandra Iliescu; Chalisa Louicharoen Cheepsunthorn; Punchalee Mungkalasut; Jean-François Trape; David Modiano; Bienvenu Sodiomon Sirima; Pratap Singhasivanon; Mark Lathrop; Anavaj Sakuntabhai; Jean-François Bureau; Philippe Gros

doi:10.1371/journal.pone.0144555

Abstract

Pyruvate kinase (PKLR) is a critical erythrocyte enzyme that is required for glycolysis and production of ATP. We have shown that Pklr deficiency in mice reduces the severity (reduced parasitemia, increased survival) of blood stage malaria induced by infection with Plasmodium chabaudi AS. Likewise, studies in human erythrocytes infected ex vivo with P. falciparum show that presence of host PK-deficiency alleles reduces infection phenotypes. We have characterized the genetic diversity of the PKLR gene, including haplotype structure and presence of rare coding variants in two populations from malaria endemic areas of Thailand and Senegal. We investigated the effect of PKLR genotypes on rich longitudinal datasets including haematological and malaria-associated phenotypes. A coding and possibly damaging variant (R41Q) was identified in the Thai population with a minor allele frequency of ~4.7%. Arginine 41 (R41) is highly conserved in the pyruvate kinase family and its substitution to Glutamine (R41Q) affects protein stability. Heterozygosity for R41Q is shown to be associated with a significant reduction in the number of attacks with Plasmodium falciparum, while correlating with an increased number of Plasmodium vivax infections. These results strongly suggest that PKLR protein variants may affect the frequency, and the intensity of malaria episodes induced by different Plasmodium parasites in humans living in areas of endemic malaria.

Citation: van Bruggen R, Gualtieri C, Iliescu A, Louicharoen Cheepsunthorn C, Mungkalasut P, Trape J-F, et al. (2015) Modulation of Malaria Phenotypes by Pyruvate Kinase (PKLR) Variants in a Thai Population. PLoS ONE 10(12): e0144555. https://doi.org/10.1371/journal.pone.0144555

Editor: Alister G. Craig, Liverpool School of Tropical Medicine, UNITED KINGDOM

Received: September 14, 2015; Accepted: November 19, 2015; Published: December 14, 2015

Copyright: © 2015 van Bruggen et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited

Data Availability: All relevant data are within the paper and its Supporting Information files.

Funding: This work was supported by research grants to PG from the Canadian Institutes for Health Research (MOP-119342) and funding to AS from the French Government’s Investissement d’Avenir program, Laboratoire d’Excellence “Integrative Biology of Emerging Infectious Diseases” (grant No. ANR-10-LABX-62-IBEID).

Competing interests: The authors have declared that no competing interests exist.

Introduction

Malaria is one of the clearest examples of host genetic contributions to susceptibility to infections (reviewed in [1–4]). Indeed, the number of clinical episodes of malaria, the level of blood parasitemia during infection, the rate of transmission (gametogenesis), and the severity of disease developed (mild, severe malaria-induced anemia, cerebral malaria), all show a strong heritable component [1, 2, 5–10]. The complexity and nature of the genetic factors regulating these traits have been studied in case-control studies with candidate genes and in a few family-based genome wide linkage analyses [1, 2, 11–14]. Genetic variants affecting invasion of erythrocytes by merozoites, intra-erythrocytic replication or elimination of parasitized RBC have a major effect on infection. For example, the Duffy antigen is the receptor for P. vivax on erythrocytes and its absence in the Duffy negative blood group prevents parasite entry in erythrocytes and protects against P. vivax malaria [12, 13]. Glycophorins (GYPA, GYPB, GYPC) bind to Plasmodium surface proteins, and GYPC-non expressing individuals show reduced invasion of erythrocytes, and are protected from infection [14]. Deletion of the anion exchanger Band 3 protein causes Melanesian ovalocytosis, which is also linked to reduced malaria incidence [15]. Heterozygosity for mutant haemoglobin (Hb) variants causing either sickle cell anemia (HbS) [16, 17] or thalassemias [4, 18–20] provide significant protection against malaria, with strong positive selection of mutant alleles in malaria-endemic areas. Glucose-6-phosphate dehydrogenase (G6PD) is required for glutathione production and protection against Hb degradation-induced oxidative stress damage. G6PD deficiency offers very strong protection against P. vivax but not against P. falciparum malaria [11]. Finally, A/B blood group antigens contribute to rosetting of parasitized RBCs, and a recent large population study has identified diminished risk of malaria in the O blood group [21, 22].

Pyruvate kinase (PK) catalyzes the last rate-limiting step of glycolysis. There are two genes in humans that code for pyruvate kinases, the liver/erythrocyte-specific enzyme (PKLR) and the muscle specific enzyme (PKM1/2). In mature erythrocytes, PKLR is essential for energy generation [23]. PKLR is active as a tetramer, removing the phosphate from phosphoenolpyruvate (PEP), and producing pyruvate and ATP [23–27]. PK-deficiency (OMIM#266200) is the most common cause of non-spherocytic hemolytic anemia, and is inherited in an autosomal recessive manner [24, 28, 29]. Clinical manifestation of PK-deficiency in humans ranges from mild compensated hemolytic anemia to severe hemolysis causing neonatal death [23, 24, 29–31]. In mice, Pklr-deficient strains AcB55 and AcB61 (Pklr^I90N) and CBA/Pk^slc (Pklr^G338D) display haemolytic anemia caused by homozygosity for loss of function mutations in the Pklr enzyme. Following infection with P. chabaudi AS, these Pklr-deficient mice show decreased peak parasitemia and decreased mortality [31, 32]. On the other hand, we have previously shown that erythrocytes from PKLR-deficient human patients are significantly less permissive to P. falciparum infection than PKLR-sufficient erythrocytes [30]. In addition, erythrocytes from heterozygous PKLR-deficient patients parasitized with P. falciparum are phagocytised at appreciably greater levels than control parasitized erythrocytes [30, 33]. Therefore, PKLR-deficiency in humans is associated ex vivo with reduced erythrocyte invasion and increased phagocytosis of early-stage infected erythrocytes.

Sequencing the PKLR gene from 387 normal individuals from different regions of the world including areas where malaria is endemic (CEPH Human Diversity Panel) showed that the Sub-Sahara African cohort exhibited the highest genetic diversity within PKLR, while the European population exhibited the lowest level of genetic diversity [34]. These findings were replicated in another study, suggesting that the PKLR gene has been under selective pressure, possibly due to malaria [35]. On the other hand, case-control studies have not detected association of PKLR variants with malarial phenotypes [35, 36]. However, disease-based cohorts with single clinical end-points (presence or absence of disease) are not optimal to test the potential protective effects of PKLR variants on human malaria. In such cohorts, it is difficult to distinguish individuals with ‘temporal’ resistance (non-infected, infected but asymptomatic, or displaying sub-clinical phenotypes at the time of sampling) from those with true resistance, whereby these individuals are infected but do not develop disease (severe or otherwise) [8]. These issues can be best addressed in longitudinal studies of community-based cohorts, wherein different quantitative malarial phenotypes such as fever, parasitemia, type and number of malaria episodes are studied over time in the same individuals. Genetic studies in these unique cohorts have increased sensitivity and can detect genetic effects on different discrete disease endophenotypes [8, 37, 38]. In this study, we took advantage of two longitudinal population studies conducted in the malaria endemic areas of Senegal and Thailand, to identify whether PKLR gene variants are associated with protection against the incidence or severity of malarial infections.

Materials and Methods

Population Studies

The population cohorts and associated longitudinal population studies in the Senegalese villages of Dielmo and Ndiop (n = 878) [7, 8, 39] and the studied Thai population (n = 897) [6, 7, 11] have been previously described. Passive and active surveillance for identification of clinical and subclinical phenotypes has been described [6–8, 11, 39–41]. Human DNA samples were obtained with written informed consent, and were approved by the ethical review Committee for Research in Human Subjects, Ministry of Public Health, Thailand. In Senegal, the project was approved by the Ministry of Health of Senegal, and by the village population. Approval is renewed on a yearly basis. Written informed consent was obtained from all participants over the age of 15 years or from the guardians of children younger than 15 year. In all cases, such consent was obtained in the presence of the school director, an independent witness. Written consent was recorded on a voluntary consent form written in both French and Wolof, the local language. This methodology was approved by the National Ethics Committee of Senegal, which carries out audits regularly along with ad hoc committees of the Ministry of Health, the Pasteur Institute (Dakar, Senegal) and the Institut de Recherche pour le Développement (Marseille, France). Patients’ records and information were de-identified prior to analysis.

Subsets of these populations were investigated for variants in PKLR, using different size exploratory and validation cohorts (see Results). Exons and exon/intron junctions of the PKLR gene were sequenced using primer pairs adapted from [34] (S1 Table). Sequences were aligned to the PKLR gene (chr1:155259084–155270792) from the hg19 (Feb 2009) human assembly (UCSC) using BioEdit Sequence Alignment Editor [42]. Haploview software [43] was used to identify haplotypes for each community using common dbSNPs (MAF≥10%). Using Custom Taqman SNP primers and probes designed for rs147659527 (L272V) and R41Q, the presence of these variants were analyzed within the Senegalese and Thai communities, respectively. In addition, we examined the R41Q variant in 340 unrelated individuals from six Southeast Asian populations (52 Mon, 62 Burmese, 65 Cambodian, 104 Thai, and 57 Lao).

Association studies and statistical analysis

The rs147659527 (L272V) genotypes were determined using custom Taqman SNP primers. The R41Q genotypes were determined using a high-resolution melting test with forward primer 5’-AAGGGCAGGTGACATGCAGT -3’ and reverse primer 5’-TGCTGCTGGAAGAAGGCAGT-3’, and analyzed with a CFX96 Touch Real-Time PCR Detection System (BIO-RAD). HRM genotypes were confirmed by PCR-RFLP, and by direct sequencing of PCR products. When possible, familial segregation of genotypes was verified using the Pedcheck software [44]. In the Thai cohort (S2 Table), we tested association of the R41Q variant with 37 blood cell counts and associated haematological parameters, as well as with 9 malaria-related phenotypes as described [6] (listed in S3 Table). All malaria-related phenotypes were analyzed for R41Q genotypes and no associations were detected except for the two phenotypes, the Plasmodium falciparum attacks and Plasmodium vivax attacks. There is no trait defining the “severity” of malaria per se in this hypo-endemic area. Attacks are defined as measured fever (axillary temperature > 37.5°C) or fever-related symptoms including headache, back pain, chills, myalgia, nausea and vomiting, associated with a blood smear positive for blood-stage trophozoite P. falciparum or P. vivax. The delay between two attacks with the same parasite is at least 30 days. For the two attack phenotypes, a Generalized Linear Mixed Model was fitted with the individual person as a factor in the random model. A Poisson error structure was implemented, thus yielding a log-linear regression. “Explanatory factors” include year (1998–2005), month, hamlet, and gender, with age factored into eight groups. The Pearson residual variance, not explained by “environmental” factors, was generated for each attack, and the sum of the residuals per person and for each parasite was used as the attack phenotype. R41Q heterozygotes were compared with R41 wild-type homozygotes for each trait using a parametric ANOVA test, taking into account normal variables affecting these traits. For non-Gaussian distribution, the non-parametric Kruskal-Wallis equality-of-populations rank test with one degree of freedom was also utilized. Analysis was completed first on non-stratified and then on gender-stratified samples. For ANOVA, mean ± SEM and the number of individuals were given for each group.

Preparation of wild-type and mutant PKLR constructs

Wild-type (WT) PKLR cDNA in a pET-21a vector was generously provided by Dr. G. Valentini (University of Pavia, Italy). An NheI restriction site (5’-G^CTAGC), Kozak consensus sequence (5’-CCACC) and an HA epitope tag (5’-TACCCATACGATGTTCCAGATTACGCT) were added to the N-terminus, while a (His)₆ tag (5’-ATGATGATGATGATGATG) was added at the C terminus via PCR amplification using primer pairs (5'-GTG CTA GCC CAC CAT GTA CCC ATA CGA TGT TCC AGA TTA CGC TTC GAT CCA GGA GAA CAT ATC ATC C -3’ and 5'-TAA TGT TCT AGA TCA ATG ATG ATG ATG ATG ATG GGA TAT GCT TAG CAC CCG CAT GA -3’). The WT construct was inserted into the pcDNA3 vector and used as template for the introduction of mutations by site-directed mutagenesis (QuikChange site-directed mutagenesis kit, Stratagene) using mutation primers (S4 Table). The veracity of mutant variants was verified by DNA sequencing (S5 Table).

Protein Expression Studies

Twenty-four hours prior to transfection, 5 x 10⁵ HEK293T cells/well were plated in complete media [DMEM (Wisent Inc., St. Bruno, QC) + 10% FBS (Wisent, Inc.) + penicillin/streptomycin (ThermoFisher Scientific, South Logan, Utah)] in 6-well tissue culture dishes. Recombinant PKLR construct DNA was mixed with Opti-MEM (Life Technologies) and Lipofectamine 2000 (Invitrogen, Carlsbad, CA) prior to being added to cells supplemented with fresh DMEM + 10% FBS. Transiently transfected cells were harvested 18 hrs later. In some experiments, transfected cells were plated in complete medium supplemented with 0.5mg/mL Geneticin (G418, Gibco Life technologies, Grand Island, NY), and G418-resistant colonies were isolated 14 days later. Transfected cells were washed in 1xPBS (HyClone, Logan, Utah), then sonicated in PBS (6x15sec) on ice (Vibra-cell Sonicator, Sonics and Materials Inc., Newtown, CT). The total protein concentration within the cell lysate was quantified using a Bradford assay (Bio-Rad Laboratories, USA), and equal amounts of protein lysate were separated on a 10% SDS-PAGE gel, followed by immunoblotting against HA.11 Clone 16B12 purified monoclonal antibody (mouse) (Covance Inc., Princeton, NJ) and β-actin antibody (mouse) (Covance Inc.) as a loading control. The secondary antibody, ECL^TM Anti-mouse IgG, Horseradish Peroxidase linked whole Ab (from sheep) (GE Healthcare Ltd., Buckinghamshire, UK), was used in conjunction with SuperSignal WestPico Chemiluminescent substrate (ThermoFisher Scientific) for protein detection.

Protein Stability Assay

For protein stability assays, cells were incubated in methionine- and cysteine-free DMEM media containing 10% dialyzed FBS, and 55 μCi of [³⁵S] methionine-cysteine (Perkin-Elmer, Boston, MA). Cells were then washed twice with PBS and incubated in chase medium (standard DMEM containing 10% FBS, 100 μg/ml methionine and 100 μg/ml cysteine) for up to 24 hours. For immunoprecipitation, equal amounts of cell lysates were incubated with the mouse anti-HA antibody overnight at 4°C, followed by incubation with protein A/G-sepharose (GE Healthcare, Piscataway, NJ) for 4 hrs. Beads were washed three times in RIPA buffer and proteins were eluted with 50μl of 3X Laemmli sample buffer. The radiolabeled proteins were separated by electrophoresis using 7.5% SDS polyacrylamide gels. The gels were then fixed, impregnated with Amplify (GE Healthcare, Piscataway, NJ), dried and exposed to film. Densitometric analysis was performed using NIH ImageJ software (NIH, Bethesda, MD).

Mice and parasite

CBA/Pk^slc mice were provided by the Japan SLC Animal Facility (Mr. H. Asai) and subsequently maintained as a breeding colony at McGill University. CBA/CaHn-Btk/J mice were purchased from Jackson Laboratories (Bar Harbor, ME, USA). All mice were kept under specific pathogen free conditions and handled according to the guidelines and regulations of the Canadian Council on Animal Care. Mice experimentation protocol was approved by the McGill Facility Animal Care Committee (P. Gros, Principal Investigator; protocol number: 5287), and include procedures to minimize distress and improve welfare. An LDH virus-free isolate of Plasmodium chabaudi AS (D. Walliker; University of Edinburgh) was maintained by weekly passage in A/J mice by intra-peritoneal infection with 10⁶ parasitized erythrocytes (pRBCs) [45]. For blood-stage infections, 8–12 weeks old mice were infected i.v. with 10⁵ pRBCs suspended in pyrogen-free PBS; blood parasitemia was monitored daily (days 4–28) on thin blood smears stained with Diff-Quik (Dade Behring). Mice infected with Plasmodium chabaudi were monitored twice daily for appearance of malaria symptoms at the peak of blood stage replication. Animals showing lethargic behavior and/or ruffled fur appearance or significant weight loss were sacrificed following anesthesia.

Results

PKLR genotypes in informative Senegalese and Thai populations

We investigated three populations living in areas of endemic malaria (Senegal; Thailand), and in which several haematological and malarial phenotypes were followed longitudinally [6–8, 11, 39]. In these populations, we tested the hypothesis that coding or non-coding variants in the PKLR gene may be associated with differential response to the malarial parasite, including episodes and intensity of infections. As a first step, we sequenced exons and intron/exon junctions of the PKLR gene in an exploratory cohort of 62 individuals from Senegal (35 from Dielmo, 27 from Ndiop) and 6 individuals from the Thai-Myanmar border region.

Nineteen single nucleotide polymorphisms (SNPs) were identified in the Senegalese communities (Table 1), including several known dbSNPs (MAF>10%) as well as a number of rarer variants. Of note, rs147659527 (c.853C>G) causes a conservative L272V substitution identified in a single homozygous Ndiop individual (Table 1). Upon further investigation, this individual did not display symptoms of PK-deficiency, suggesting that L272V does not affect PKLR protein activity and/or is phenotypically neutral. PKLR haplotypes were defined using eight SNPs with MAF>10% (rs8177964, rs3020781, rs2071053, rs4620533, rs1052176, rs1052177, rs932972, and rs8847). The Dielmo and Ndiop communities displayed five haplotypes each, three of which were shared between them, with the remaining less frequent haplotypes being unique to each community (Fig 1).

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Fig 1. PKLR gene variants discovered in both the Senegalese and Thai community.

(a) Location of coding and non-coding variants within the PKLR gene. Encoded on chromosome 1q22 (155259084–155270792 base pairs), the erythrocyte isoform, transcribed from a tissue specific promoter has 11 exons (blocks). (b) Haplotypes and haplotype frequencies for sequenced population. Determined using the Haploview software, haplotypes were defined by the SNPs with a MAF ≥ 10% that defined the Senegalese and Thai populations. Seven of the SNPs were shared within the three study populations, while rs4620533 was specific for the Senegalese populations and rs3762272 was found only within the Thai population. Haplotype frequency distribution were calculated for the complete data set (ALL), or subdivided on country (Thailand, THA; Senegal, SEN) or village (Dielmo, DLM; Ndiop, NDP).

https://doi.org/10.1371/journal.pone.0144555.g001

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Table 1. Single nucleotide polymorphisms and alternative allele frequencies within the sequenced population.

https://doi.org/10.1371/journal.pone.0144555.t001

In the six Thai individuals sequenced, several common dbSNPs were present in addition to a novel non-synonymous mutation (c.161A>G) detected in a single individual (Table 1), resulting in a coding change from an arginine at residue 41 to glutamine (R41Q) (Fig 2). R41 is highly conserved across species and the arginine to glutamine substitution is not conservative and results in the loss of the positive side chain of wild-type R41. Sequencing an expanded cohort of 96 Thai individuals identified nine samples positive for R41Q (all heterozygotes), establishing an allele frequency of ~5% for this coding variant in the Thai population. Eleven additional variants with minor allele frequency > 1% were detected in the Thai cohort, including rs200695047 (c.844G>T) which codes for a valine to phenylalanine substitution at amino acid 269 (V269F). This mutation was identified in one homozygous and six heterozygous samples. V269 is not conserved throughout evolution (Fig 2) and the V269F homozygote did not display symptoms of PK-deficiency, suggesting that V269F does not affect PKLR protein activity. Haplotype analysis with informative SNPs (rs3020781, rs2071053, rs4620533, rs3762272, rs1052176, rs1052177, rs932972, rs8847) revealed 3 haplotypes in the Thai sample, with the most common haplotype (68.9%) being unique to the Thai community, and the two less frequent haplotypes being shared with the Senegalese samples (Fig 1). Interestingly, the R41Q variant was detected on the Thai-specific haplotype (Haplotype 1, Fig 1) suggesting that this mutation may be population-specific, having arisen on a Southeast Asian genetic background.

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Fig 2. PKLR coding variants identified in the test populations.

(a) Schematic representation of the PKLR protein, location of catalytically active residues (phosphoenolpyruvate, PEP, binding residues; R116, G338, D339, and T371) and allosteric regulation site (fructose-1,6-bisphosphate, FBP, binding site; loops 475–490 and 557–566, and residue R532) within structural domains [34, 46], and location of non-synonymous mutations identified from sequencing (R41Q, V269F and L272V). (b) Subset of multiple species protein alignment by Clustal Omega (www.ebi.ac.uk) where “*” = single, fully conserved residue; “:” = strongly similar properties

https://doi.org/10.1371/journal.pone.0144555.g002

Population genetic analysis of the R41Q and L272V variants in PKLR

Further characterization of possible associations between the L272V and R41Q variants and malarial phenotypes was undertaken, as these variants affect residues that are highly conserved in the PKLR protein family and may affect PKLR function. The L272V mutation (rs147659527) was identified in one homozygous individual from the Senegalese community of Ndiop. Further genotyping of the Ndiop community revealed that L272V was present in a single family, where the grandmother was homozygous, and three of her children and a grandchild were heterozygous. This family is one of 12 Fulani families in the Ndiop community (85 individuals), an ethnic group distributed throughout Western and Central Africa [18, 47, 48]. Additional sequencing of the PKLR gene in 149 unrelated individuals of Mossi (n = 89) and Fulani (n = 60) ethnicity from Burkina Faso identified three L272V heterozygotes in Mossi individuals. Hence, the L272V variant appears to be a rare variant (0.2%-1.5%; dbSNP build 138) distributed in different African populations. The relatively rarity of this variant in the African populations available was too low to examine possible association with haematological or malarial phenotypes.

Amongst the 102 Thai samples sequenced, we detected ten R41Q heterozygotes. Genotyping this mutation in the entire study population consisting of 871 samples from 181 families, and from 46 unrelated individuals identified one homozygote individual and 81 heterozygotes (71 of Karen origin), representing a mutant allele frequency of approximately 4.7% in this population. The R41Q genotypes exist in Hardy-Weinberg equilibrium (HWE) in this Thai sample set. The homozygous R41Q mutant individual did not exhibit extreme blood phenotypes, although its erythrocyte and reticulocyte counts were both low (falling within the lowest 10^th and 25^th percentile; data not shown). The predicted allele frequency of 5% may provide sufficient statistical power for association studies.

Genotyping for the R41Q mutation in an additional 340 unrelated Southeast Asian control samples, detected 13 R41Q heterozygotes, consisting of 2 Cambodians, 2 Laos, 2 Thais, 4 Myanmarese, and 3 Mons. The R41Q genotypes exist in HWE in this sample set with the allele frequency in each population, 1.5% in Cambodians, 1.8% in Laos, 1% in Thais (mixed ethnicity), 3.2% in Myanmarese, and 2.9% in Mon. It is interesting to note that the frequency of the R41Q allele is low in the general Thai population (~1%), while it is higher in the Thais of Karen ethnicity (~5%) who generally live in malaria-endemic areas. Similarly, the frequency of the R41Q allele is higher in the Myanmarese population (~3.2%), where a large population of ethnic Karen reside [49].

Association studies

Among the 871 Thai individuals genotyped for R41Q, one individual was homozygous for the mutation, 81 were heterozygous and 789 had the wild-type genotype. Associations between R41Q heterozygosity and various blood and malarial parameters listed in S3 Table and previously described [6], were examined. No association was detected between R41Q status and erythrocyte count, hematocrit, or reticulocyte levels and any other phenotypes except two. Indeed, a significant statistical association was noted between R41Q heterozygosity and a reduced frequency of Plasmodium falciparum attacks (mean frequency of heterozygotes (m_He) = -0.23 ± 0.09 (n = 65) and mean frequency of homozygous wild-type (m_Ho) = 0.01 ± 0.03 (n = 649); ANOVA p = 0.0097, Kruskal-Wallis (KW) p = 0.0022), with this association being significant only in females (m_He = -0.30 ± 0.11 (n = 34) and m_Ho = 0.01 ± 0.05 (n = 346); ANOVA p = 0.0260, KW p = 0.0053 versus m_He = -0.16 ± 0.13 (n = 31) and m_Ho = 0.02 ± 0.05 (n = 303) ANOVA p = 0.17, KW p = 0.1436 for males) (Fig 3A). Additionally, we determined that individuals heterozygous for R41Q are significantly more susceptible to Plasmodium vivax attacks than Plasmodium falciparum attacks (m_He = 0.49 ± 0.18 (n = 65) and m_Ho = -0.07 ± 0.04 (n = 649); ANOVA p = 1.32 10⁻⁴) again limited to females (m_He = 0.84 ± 0.28 (n = 34) and m_Ho = -0.07 ± 0.05 (n = 346), ANOVA p = 7.1 10⁻⁵) and not males (m_He = 0.10 ± 0.15 (n = 31) and m_Ho = -0.08 ± 0.06 (n = 303), ANOVA p = 0.31). This result strongly suggests that females and not males are differently susceptible to Plasmodium falciparum attacks and to Plasmodium vivax attacks depending on R41Q heterozygosity. Gender-specific effects for blood-stage malaria phenotypes have been previously reported in humans [11], and mice [45]. These findings suggest that heterozygosity for R41Q may have an impact on host susceptibility to infection with malarial parasites in endemic areas of disease.

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Fig 3. Effect of human and mouse PKLR variants on malaria phenotypes.

(a) Association of the R41Q mutation and Plasmodium falciparum attacks in the genotyped Thai population. Residual Plasmodium falciparum attacks for heterozygous (R41Q) and homozygous wild-type (R41R) individuals. Both parametric and non-parametric tests indicate statistically significant decrease in attacks for heterozygous females only (ANOVA p = 0.0260 Kruskal-Wallis (KW) p = 0.0053). (b) Residual Plasmodium vivax attacks for heterozygous (R41Q) and homozygous wild-type (R41R) individuals. A statistically significant increase in attacks was noted in heterozygous females though only for the parametric test (ANOVA p = 0.0007), not the non-parametric test (KW p = 0.1789). (c) Time course of P. chabaudi chabaudi AS infection in PK-sufficient (CBA/CaHn-Btk^xid/J, circles), PK-deficient (CBA/Pk^slc, triangles) and F1 generation mice (squares). (d) Peak parasitemia (8dpi). Results are separated based on gender. Statistical significance was determined via unpaired, two-tailed t-tests, calculated by GraphPad (Prism) where “*” represents p<0.05. Statistically significant differences between PK-sufficient and F1 heterozygotes was noted for both genders (females: p = 0.0264, males: p = 0.0378).

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Heterozygosity for Pklr in mice protects against peak parasitemia

In humans and mice, pyruvate kinase deficiency is inherited in a strictly recessive fashion, with heterozygotes displaying no clinical symptoms, and displaying normal haematological profiles. Nevertheless, we have previously shown that P. falciparum-parasitized erythrocytes from subjects heterozygous for loss of function at PKLR are phagocytized more avidly than similarly infected wild-type erythrocytes [30, 33]. Hence, we tested the effect of heterozygosity for the known loss of function mutation at murine Pklr (G338D) [50], on haematological replication of the malarial parasite in a mouse model of infection with P. chabaudi. For this, wild-type controls (CBA/CaHn-Btk^xid/J), pyruvate kinase deficient mutants (CBA/Pk^slc) homozygous for a loss of function allele at Pklr (G338D), as well as F1 heterozygotes (G338D/+) were infected with P. chabaudi AS and blood parasitemia was followed over time (Fig 3C). We observed that male mice of all genotypes were more susceptible than female mice, in agreement with a previous report [45]. However, we noted that at the peak of infection (peak parasitemia), G338D/+ heterozygotes displayed lower peak parasitemia values than their wild-type counterparts (CBA/J), and this for both male (two-tailed unpaired t-test p = 0.0378) and female mice (p = 0.0264) (Fig 3D). In males, this significant difference in blood parasitemia between wild-type and G338/+ heterozygotes extended from day 7–9 post-infection (p = 0.0188; p = 0.0290). This suggests that heterozygosity for mutant Pklr variant in mice improves response to infection with the malarial parasite.

R41Q recombinant protein exhibits modest effect on half-life

Overexpression of the R41Q variant in transiently transfected HEK293T cells, followed by semi-quantitative measurement of pyruvate kinase activity (PK-LDH assay) failed to detect a major effect of the R41Q mutation on basal enzymatic activity in such over-expressing cells (data not shown). We also examined the effect of the R41Q mutation on stability of the transiently transfected [³⁵S]-methionine labelled protein in pulse-chase studies (Fig 4). In three independent experiments, we observed a shorter half-life for R41Q compared to WT protein expressed in transfected cells: The effect was modest but reproducible, suggesting that R41Q may affect protein stability in these cells.

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Fig 4. Effect of R41Q on PKLR protein stability.

(a) Transiently transfected HEK293T cells expressing either the wild-type (WT) or the R41Q mutant PKLR proteins were metabolically labelled with [³⁵S]-methionine followed by chase in label-free media for the indicated time period (in hours). Cell lysates were prepared and PKLR was immunoprecipitated, followed by gel electrophoresis. (b) The radiolabelled PKLR species were quantitated using ImageJ software, and averages from 2–3 independent experiments were calculated as a fraction of the maximum labelling observed at 0 hr.

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Discussion

Malaria kills one child every minute in Sub-Saharan Africa [51, 52]. Over time, selective pressure from such a lethal infection has resulted in the genetic retention in malaria endemic areas of certain allelic variants that confer partial protection against disease. The erythrocyte is the major replication niche of the Plasmodium parasite, and variants in certain erythrocyte proteins have been found enriched in malaria-endemic areas. Although homozygosity for such variants causes disease on their own (sickle cell anemia, thalassemias, G6PD deficiency), heterozygosity confers various degrees of protection against malaria [1–3, 53, 54]. We have previously shown that homozygosity for loss of function mutations within the erythrocyte pyruvate kinase gene (Pklr) in mice reduces the severity of infection with the blood stage parasite P. chabaudi AS [31, 32]. Likewise, in human erythrocytes infected ex vivo with P. falciparum, homozygosity for PK-deficiency alleles causes a reduction in parasite replication, while parasitized erythrocytes from heterozygotes are phagocytized more avidly than those from normal individuals [30, 33]. Using comparative analyses of archival populations contained in the CEPH Human Diversity panel from current and ancestral areas of malaria, we detected rich genetic diversity in the PKLR gene, including the presence of rare, potential loss-of-function protein variants. Neutrality testing of PKLR haplotypes and variants suggested positive selection of the gene in the sub-Saharan African and Pakistani populations, possibly by the malarial parasite [34]. This hypothesis remained to be formally tested, and was the major aim of the current study.

Herein, we probed for the possible association between rare or common variants at the sequenced PKLR gene, and haematological and malaria-associated phenotypes in three populations from Thailand and Senegal, living in areas of seasonal or year-long exposure to the malarial parasite. These populations were studied longitudinally for multiple phenotypes, including but not limited to frequency and intensity of infection by the malarial parasites, thereby providing added power to detect genetic effects of the host in response to the parasite. The Thai population has been used in the past to document the effect of G6PD deficiency (Mahidol allele) on differential susceptibility to infection with P. falciparum and P. vivax [11].

Sequencing the PKLR gene in subsets of Senegalese and Thai individuals identified a number of common variants (SNPs), nine of which were used to define PKLR haplotypes in these populations. Eight PKLR haplotypes were detected, two of which were common and representing approximately 86% of the Senegalese and 31% of the Thai populations. These common haplotypes are also present in various North, South, and Sub-Saharan African, Pakistani, Southeast Asian and European populations [34]. Haplotype analysis at PKLR further detected population substructure in the two Senegalese populations. The Dielmo community is composed primarily of the Serere and Mandinka ethnic groups [7], while the Ndiop community is comprised predominantly of Wolof and Fulani ethnicities [7]: unique PKLR haplotypes were detected in these populations. Of particular interest is the Thai-specific haplotype 1 (Fig 1) onto which the R41Q mutation appeared to have emerged. This strongly suggests that the R41Q variant is population specific. We also detected the R41Q variant in Cambodian, Lao, Myanmarese, and Mon populations from other malaria-endemic areas. Although malaria-associated phenotypes are not available for these populations, presence of the R41Q mutation suggests that the variant occurred on an ancient South-Asian haplotype which is shared amongst populations of Thailand, Laos, and Cambodia.

Sequencing also identified three coding variants in the PKLR protein; these variants affected residues that were either highly conserved (R41, L272), or coded for a very significant change in amino acid property (V269F), both of which possibly result in altered protein function. The L272V and V269F variants were present at very low frequency in the two populations, and association with haematological and malaria traits could not be conducted with sufficient statistical power. Genotyping revealed that the R41Q mutation was present with a minor allele frequency of approximately 4.5% within the Thai cohort. We noted a statistically significant association between R41Q heterozygosity and a reduced frequency of Plasmodium falciparum attacks, with this association being significant only in females. Additionally, there appears to be a tendency for R41Q heterozygotes to be more susceptible to infection with Plasmodium vivax, although this was only significant in parametric, rather than non-parametric tests, and again limited to females (Fig 3A and 3B). Although these findings require replication in additional cohorts of individuals with informative R41Q genotypes and from malaria-endemic areas, the current results suggest a possible association of the R41Q variant with reduced susceptibility to P. falciparum infection. The increased susceptibility to P. vivax infection noted in some R41Q heterozygotes and concomitant to reduced susceptibility to P. falciparum may reflect immunological cross-reactivity and immune protection in regions of mixed infections [55]. On the other hand, such genetically influenced differential susceptibility to different malarial parasites has been previously reported in the case of glucose-6-phosphate dehydrogenase deficiency. Indeed, it was demonstrated that the G6PD^Mahidol (487A) variant reduces infection with P. vivax, but not with P. falciparum in a population from South Asia, suggesting that Plasmodium vivax has been a driving force behind the strong selective advantage conferred by this mutation [11].

Pyruvate kinase is composed of an amino terminal (N) domain with a helix-turn-helix motif, followed by a bi-partite A domain (A1, A2) that contains residues implicated in binding the phosphoenolpyruvate (PEP) substrate, as well as a conserved B domain linker, followed by the C-terminal domain that binds the allosteric regulator fructose-1,6-bisphosphate (FBP) (Fig 2). Structure function studies have shown that PKLR exists either in the active R-state or the inactive T-state, and that the state of activation is regulated by intra- and inter-domain interactions between the A and C domains of adjacent subunits as well as the A/B and A/C domains within a subunit [26, 56, 57]. The R41Q mutation is located within the N-terminal domain of PKLR, and the arginine residue is highly conserved in mammals. While overexpression of the R41Q variant in transfected cells failed to identify a significant effect of the mutation on enzymatic activity, we noted a modest but reproducible effect of R41Q on protein half-life in pulse-chase studies.

In addition to basal activity, there are number of structural and functional aspects of PKLR that may be affected by R41Q. PKLR activity is regulated in part by phosphorylation of serine 43, which causes increased affinity for allosteric inhibitors ATP and alanine, and decreased affinity for PEP and the allosteric activator FBP [58, 59]. Two mutations mapping to the N-terminal domain of PKLR (near R41Q) have been functionally characterized. One mutation (A36G) identified in a case of congenital nonspherocytic hemolytic anemia with PK-deficiency, was shown to partially impair enzyme activity [60]. A possible gain-of-function variant (G37E) was found associated with enzyme hyper-activity, increased ATP levels, and possibly linked to altered phosphorylation of S43 [61, 62]. Additionally, the PK-R isoform undergoes post-translational proteolytic cleavage of the N-terminus during erythrocyte maturation, generating full length and truncated proteins with different capacity for homo- and heterotetramerization [24, 26, 46, 57, 63]. Also, fructose-1,6-bisphosphate promotes tetramer formation, while ATP causes tetramer disassociation (PK-R/L, M2) [64, 65]. Together, these findings suggest that the N-terminal domain of PKLR that carries the R41Q variant plays a key role in enzyme function, regulation of activity and functional oligomerization. Finally, PKLR monomers can oligomerize with monomers from the other isoform, PK-M2, leading to the formation of hybrids that contain 1 to 3 subunits of either isoform [66–69]. PK-M2 is broadly expressed in most cells lines used in transfection studies, including HEK-293 cells used herein, which further complicates the functional study of PKLR variants expressed in these cells. Indeed, it is possible that R41Q affects the ability of the PK-R enzyme to heterodimerize with background PK-M2 proteins present in transfected cells, leading to PK multimers of unknown activity.

Finally, it has been observed that some disease-associated PKLR mutations do not affect enzymatic activity of the purified protein. For example, the R510Q mutant in a patient with nonspherocytic hemolytic anemia shows wild-type kinetic activity and regulatory properties when expressed in bacteria [57]: However, R510Q exhibits significantly lower thermal stability when compared to the wild-type control, suggesting that protein instability rather than reduced enzymatic activity is responsible for PK-deficiency in R510Q patients [57]. Additional studies will be required to characterize the molecular effect of the R41Q variant on PKLR structure and function.

Although our study requires additional validation in other cohorts where longitudinal data on haematological malaria phenotypes are available, results obtained so far strongly suggest that allelic variants at PKLR modulate susceptibility to Plasmodium infection in humans, with protective alleles possibly retained in populations living in areas of endemic malaria. The cumulative data supporting this proposal include: a) the demonstration that homozygosity for loss of function at Pklr cause complete protection against blood-stage malaria (P. chabaudi AS) in mice [31, 32], with heterozygotes showing reduced levels of blood-stage parasitemia (Fig 3); b) homozygosity and heterozygosity for human PK-deficiency associated loss of function alleles at PKLR cause a reduction in parasite replication and increase phagocytosis of parasitized erythrocytes in studies with primary erythrocytes infected ex vivo with P. falciparum [30]; c) the PKLR gene shows highest sequence diversity in regions where malaria is or has been endemic, when compared to regions where it has not been prevalent [34]; d) a unique PKLR variant (R41Q) with reduced protein stability has appeared on a common haplotype from Southeast Asia and shows an association with reduced number of malaria attacks in females from a large Thai cohort of Karen ethnicity from a malaria endemic area. Therefore, along with haemoglobin, glucose-6-phosphate dehydrogenase, anion exchanger band 3, and several others, PKLR would represent another example of an erythrocyte-specific protein in which heterozygosity for partial or complete loss of function variants confer some degree of protection against malaria.

Supporting Information

S1 Table. Sequencing primers designed to span the exon and intron/exon junctions of the PKLR gene.

Adapted from Berghout et. al. [34].

https://doi.org/10.1371/journal.pone.0144555.s001

(DOCX)

S2 Table. Main characteristic of the Thai cohort (n = 897) investigated in this study.

https://doi.org/10.1371/journal.pone.0144555.s002

(DOCX)

S3 Table. List of haematological and malaria phenotypes investigated in this study.

https://doi.org/10.1371/journal.pone.0144555.s003

(DOCX)

S4 Table. Site directed mutagenesis primers.

Primers were designed to have at least 10 nucleotides flanking the single point mutation (in bold).

https://doi.org/10.1371/journal.pone.0144555.s004

(DOCX)

S5 Table. Construct sequencing primers.

Designed within the PKLR cDNA insert and pcDNA3 vector, these primers were utilized in confirming the sequence of the wild-type and mutant constructs.

https://doi.org/10.1371/journal.pone.0144555.s005

(DOCX)

Acknowledgments

The authors are indebted to Normand Groulx and Susan Gauthier for expert technical assistance.

Author Contributions

Conceived and designed the experiments: ML AS JFB PG. Performed the experiments: RvB CG AI. Analyzed the data: RvB CG JFB. Contributed reagents/materials/analysis tools: CLC PM JFT DM BSS PS. Wrote the paper: RvB PG.

References

1. Kwiatkowski DP. How malaria has affected the human genome and what human genetics can teach us about malaria. Am J Hum Genet. 2005;77(2):171–92. pmid:16001361
- View Article
- PubMed/NCBI
- Google Scholar
2. Bongfen SE, Laroque A, Berghout J, Gros P. Genetic and genomic analyses of host-pathogen interactions in malaria. Trends Parasitol. 2009;25(9):417–22. pmid:19717339
- View Article
- PubMed/NCBI
- Google Scholar
3. Min-Oo G, Gros P. Erythrocyte variants and the nature of their malaria protective effect. Cell Microbiol. 2005;7(6):753–63. pmid:15888079
- View Article
- PubMed/NCBI
- Google Scholar
4. Williams TN, Wambua S, Uyoga S, Macharia A, Mwacharo JK, Newton CR, et al. Both heterozygous and homozygous alpha+ thalassemias protect against severe and fatal Plasmodium falciparum malaria on the coast of Kenya. Blood. 2005;106(1):368–71. pmid:15769889
- View Article
- PubMed/NCBI
- Google Scholar
5. Rihet P, Traore Y, Abel L, Aucan C, Traore-Leroux T, Fumoux F. Malaria in humans: Plasmodium falciparum blood infection levels are linked to chromosome 5q31-q33. Am J Hum Genet. 1998;63(2):498–505. pmid:9683598
- View Article
- PubMed/NCBI
- Google Scholar
6. Phimpraphi W, Paul R, Witoonpanich B, Turbpaiboon C, Peerapittayamongkol C, Louicharoen C, et al. Heritability of P. falciparum and P. vivax malaria in a Karen population in Thailand. PLoS One. 2008;3(12):e3887. pmid:19060954
- View Article
- PubMed/NCBI
- Google Scholar
7. Lawaly YR, Sakuntabhai A, Marrama L, Konate L, Phimpraphi W, Sokhna C, et al. Heritability of the human infectious reservoir of malaria parasites. PLoS One. 2010;5(6):e11358. pmid:20613877
- View Article
- PubMed/NCBI
- Google Scholar
8. Sakuntabhai A, Ndiaye R, Casademont I, Peerapittayamongkol C, Rogier C, Tortevoye P, et al. Genetic determination and linkage mapping of Plasmodium falciparum malaria related traits in Senegal. PLoS One. 2008;3(4):e2000. pmid:18431485
- View Article
- PubMed/NCBI
- Google Scholar
9. Timmann C, Evans JA, Konig IR, Kleensang A, Ruschendorf F, Lenzen J, et al. Genome-wide linkage analysis of malaria infection intensity and mild disease. PLoS Genet. 2007;3(3):e48. pmid:17381244
- View Article
- PubMed/NCBI
- Google Scholar
10. Jallow M, Teo YY, Small KS, Rockett KA, Deloukas P, Clark TG, et al. Genome-wide and fine-resolution association analysis of malaria in West Africa. Nat Genet. 2009;41(6):657–65. pmid:19465909
- View Article
- PubMed/NCBI
- Google Scholar
11. Louicharoen C, Patin E, Paul R, Nuchprayoon I, Witoonpanich B, Peerapittayamongkol C, et al. Positively selected G6PD-Mahidol mutation reduces Plasmodium vivax density in Southeast Asians. Science. 2009;326(5959):1546–9. pmid:20007901
- View Article
- PubMed/NCBI
- Google Scholar
12. Zimmerman PA, Woolley I, Masinde GL, Miller SM, McNamara DT, Hazlett F, et al. Emergence of FY*A(null) in a Plasmodium vivax-endemic region of Papua New Guinea. Proc Natl Acad Sci U S A. 1999;96(24):13973–7. pmid:10570183
- View Article
- PubMed/NCBI
- Google Scholar
13. Miller LH, Mason SJ, Clyde DF, McGinniss MH. The resistance factor to Plasmodium vivax in blacks. The Duffy-blood-group genotype, FyFy. N Engl J Med. 1976;295(6):302–4. pmid:778616
- View Article
- PubMed/NCBI
- Google Scholar
14. Maier AG, Duraisingh MT, Reeder JC, Patel SS, Kazura JW, Zimmerman PA, et al. Plasmodium falciparum erythrocyte invasion through glycophorin C and selection for Gerbich negativity in human populations. Nat Med. 2003;9(1):87–92. pmid:12469115
- View Article
- PubMed/NCBI
- Google Scholar
15. Genton B, al-Yaman F, Mgone CS, Alexander N, Paniu MM, Alpers MP, et al. Ovalocytosis and cerebral malaria. Nature. 1995;378(6557):564–5. pmid:8524388
- View Article
- PubMed/NCBI
- Google Scholar
16. Friedman MJ. Erythrocytic mechanism of sickle cell resistance to malaria. Proc Natl Acad Sci U S A. 1978;75(4):1994–7. pmid:347452
- View Article
- PubMed/NCBI
- Google Scholar
17. Ayi K, Turrini F, Piga A, Arese P. Enhanced phagocytosis of ring-parasitized mutant erythrocytes: a common mechanism that may explain protection against falciparum malaria in sickle trait and beta-thalassemia trait. Blood. 2004;104(10):3364–71. pmid:15280204
- View Article
- PubMed/NCBI
- Google Scholar
18. Modiano D, Luoni G, Sirima BS, Lanfrancotti A, Petrarca V, Cruciani F, et al. The lower susceptibility to Plasmodium falciparum malaria of Fulani of Burkina Faso (west Africa) is associated with low frequencies of classic malaria-resistance genes. Trans R Soc Trop Med Hyg. 2001;95(2):149–52. pmid:11355545
- View Article
- PubMed/NCBI
- Google Scholar
19. Olivieri NF, Muraca GM, O'Donnell A, Premawardhena A, Fisher C, Weatherall DJ. Studies in haemoglobin E beta-thalassaemia. Br J Haematol. 2008;141(3):388–97. pmid:18410572
- View Article
- PubMed/NCBI
- Google Scholar
20. Mockenhaupt FP, Ehrhardt S, Gellert S, Otchwemah RN, Dietz E, Anemana SD, et al. Alpha(+)-thalassemia protects African children from severe malaria. Blood. 2004;104(7):2003–6. pmid:15198952
- View Article
- PubMed/NCBI
- Google Scholar
21. Rowe JA, Opi DH, Williams TN. Blood groups and malaria: fresh insights into pathogenesis and identification of targets for intervention. Curr Opin Hematol. 2009;16(6):480–7. pmid:19812491
- View Article
- PubMed/NCBI
- Google Scholar
22. Fry AE, Griffiths MJ, Auburn S, Diakite M, Forton JT, Green A, et al. Common variation in the ABO glycosyltransferase is associated with susceptibility to severe Plasmodium falciparum malaria. Hum Mol Genet. 2008;17(4):567–76. pmid:18003641
- View Article
- PubMed/NCBI
- Google Scholar
23. van Wijk R, van Solinge WW. The energy-less red blood cell is lost: erythrocyte enzyme abnormalities of glycolysis. Blood. 2005;106(13):4034–42. pmid:16051738
- View Article
- PubMed/NCBI
- Google Scholar
24. Zanella A, Bianchi P. Red cell pyruvate kinase deficiency: from genetics to clinical manifestations. Baillieres Best Pract Res Clin Haematol. 2000;13(1):57–81. pmid:10916678
- View Article
- PubMed/NCBI
- Google Scholar
25. Kanno H, Fujii H, Miwa S. Structural analysis of human pyruvate kinase L-gene and identification of the promoter activity in erythroid cells. Biochem Biophys Res Commun. 1992;188(2):516–23. pmid:1445295
- View Article
- PubMed/NCBI
- Google Scholar
26. van Wijk R, Huizinga EG, van Wesel AC, van Oirschot BA, Hadders MA, van Solinge WW. Fifteen novel mutations in PKLR associated with pyruvate kinase (PK) deficiency: structural implications of amino acid substitutions in PK. Hum Mutat. 2009;30(3):446–53. pmid:19085939
- View Article
- PubMed/NCBI
- Google Scholar
27. Gupta RK, Oesterling RM. Dual divalent cation requirement for activation of pyruvate kinase; essential roles of both enzyme- and nucleotide-bound metal ions. Biochemistry. 1976;15(13):2881–7. pmid:7293
- View Article
- PubMed/NCBI
- Google Scholar
28. Valentine WN, Tanaka KR, Miwa S. A specific erythrocyte glycolytic enzyme defect (pyruvate kinase) in three subjects with congenital non-spherocytic hemolytic anemia. Trans Assoc Am Physicians. 1961;74:100–10. pmid:13924348
- View Article
- PubMed/NCBI
- Google Scholar
29. Hirono A, Kanno H, Miwa S, Beutler E. Pyruvate kinase deficiency and other enzymopathies of the erythrocyte. In: Scriver CR, editor. The Metabolic and Molecular Bases of Inherited Disease. 8th ed ed. New York: McGraw-Hill; 2001. p. 4637–64.
30. Ayi K, Min-Oo G, Serghides L, Crockett M, Kirby-Allen M, Quirt I, et al. Pyruvate kinase deficiency and malaria. N Engl J Med. 2008;358(17):1805–10. pmid:18420493
- View Article
- PubMed/NCBI
- Google Scholar
31. Min-Oo G, Fortin A, Tam MF, Gros P, Stevenson MM. Phenotypic expression of pyruvate kinase deficiency and protection against malaria in a mouse model. Genes Immun. 2004;5(3):168–75. pmid:15029238
- View Article
- PubMed/NCBI
- Google Scholar
32. Min-Oo G, Fortin A, Tam MF, Nantel A, Stevenson MM, Gros P. Pyruvate kinase deficiency in mice protects against malaria. Nat Genet. 2003;35(4):357–62. pmid:14595440
- View Article
- PubMed/NCBI
- Google Scholar
33. Durand PM, Coetzer TL. Pyruvate kinase deficiency protects against malaria in humans. Haematologica. 2008;93(6):939–40. pmid:18460648
- View Article
- PubMed/NCBI
- Google Scholar
34. Berghout J, Higgins S, Loucoubar C, Sakuntabhai A, Kain KC, Gros P. Genetic diversity in human erythrocyte pyruvate kinase. Genes Immun. 2012;13(1):98–102. pmid:21833022
- View Article
- PubMed/NCBI
- Google Scholar
35. Machado P, Manco L, Gomes C, Mendes C, Fernandes N, Salome G, et al. Pyruvate kinase deficiency in sub-Saharan Africa: identification of a highly frequent missense mutation (G829A;Glu277Lys) and association with malaria. PLoS One. 2012;7(10):e47071. pmid:23082140
- View Article
- PubMed/NCBI
- Google Scholar
36. Alves J, Machado P, Silva J, Goncalves N, Ribeiro L, Faustino P, et al. Analysis of malaria associated genetic traits in Cabo Verde, a melting pot of European and sub Saharan settlers. Blood Cells Mol Dis. 2010;44(1):62–8. pmid:19837619
- View Article
- PubMed/NCBI
- Google Scholar
37. Phimpraphi W, Paul RE, Yimsamran S, Puangsa-art S, Thanyavanich N, Maneeboonyang W, et al. Longitudinal study of Plasmodium falciparum and Plasmodium vivax in a Karen population in Thailand. Malar J. 2008;7:99. pmid:18518964
- View Article
- PubMed/NCBI
- Google Scholar
38. Trape JF, Rogier C, Konate L, Diagne N, Bouganali H, Canque B, et al. The Dielmo project: a longitudinal study of natural malaria infection and the mechanisms of protective immunity in a community living in a holoendemic area of Senegal. Am J Trop Med Hyg. 1994;51(2):123–37. pmid:8074247
- View Article
- PubMed/NCBI
- Google Scholar
39. Rogier C, Commenges D, Trape JF. Evidence for an age-dependent pyrogenic threshold of Plasmodium falciparum parasitemia in highly endemic populations. Am J Trop Med Hyg. 1996;54(6):613–9. pmid:8686780
- View Article
- PubMed/NCBI
- Google Scholar
40. Rogier C, Ly AB, Tall A, Cisse B, Trape JF. Plasmodium falciparum clinical malaria in Dielmo, a holoendemic area in Senegal: no influence of acquired immunity on initial symptomatology and severity of malaria attacks. Am J Trop Med Hyg. 1999;60(3):410–20. pmid:10466970
- View Article
- PubMed/NCBI
- Google Scholar
41. Rogier C, Tall A, Diagne N, Fontenille D, Spiegel A, Trape JF. Plasmodium falciparum clinical malaria: lessons from longitudinal studies in Senegal. Parassitologia. 1999;41(1–3):255–9. pmid:10697865
- View Article
- PubMed/NCBI
- Google Scholar
42. Hall TA. BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symposium Series. 1999;41:95–8.
- View Article
- Google Scholar
43. Barrett JC, Fry B, Maller J, Daly MJ. Haploview: analysis and visualization of LD and haplotype maps. Bioinformatics. 2005;21(2):263–5. pmid:15297300
- View Article
- PubMed/NCBI
- Google Scholar
44. O'Connell JR, Weeks DE. PedCheck: a program for identification of genotype incompatibilities in linkage analysis. Am J Hum Genet. 1998;63(1):259–66. pmid:9634505
- View Article
- PubMed/NCBI
- Google Scholar
45. Fortin A, Cardon LR, Tam M, Skamene E, Stevenson MM, Gros P. Identification of a new malaria susceptibility locus (Char4) in recombinant congenic strains of mice. Proc Natl Acad Sci U S A. 2001;98(19):10793–8. pmid:11535821
- View Article
- PubMed/NCBI
- Google Scholar
46. Valentini G, Chiarelli LR, Fortin R, Dolzan M, Galizzi A, Abraham DJ, et al. Structure and function of human erythrocyte pyruvate kinase. Molecular basis of nonspherocytic hemolytic anemia. J Biol Chem. 2002;277(26):23807–14. pmid:11960989
- View Article
- PubMed/NCBI
- Google Scholar
47. Modiano D, Petrarca V, Sirima BS, Bosman A, Nebie I, Diallo D, et al. Plasmodium falciparum malaria in sympatric ethnic groups of Burkina Faso, west Africa. Parassitologia. 1995;37(2–3):255–9. pmid:8778668
- View Article
- PubMed/NCBI
- Google Scholar
48. Tiono AB, Sirima SB, Hamed K. Fulani show decreased susceptibility to Plasmodium falciparum infection versus Mossi: data from a community-wide screening and treatment of asymptomatic carriers in Burkina Faso. Malar J. 2013;12:163. pmid:23680454
- View Article
- PubMed/NCBI
- Google Scholar
49. Summerer M, Horst J, Erhart G, Weissensteiner H, Schonherr S, Pacher D, et al. Large-scale mitochondrial DNA analysis in Southeast Asia reveals evolutionary effects of cultural isolation in the multi-ethnic population of Myanmar. BMC Evol Biol. 2014;14:17. pmid:24467713
- View Article
- PubMed/NCBI
- Google Scholar
50. Morimoto M, Kanno H, Asai H, Tsujimura T, Fujii H, Moriyama Y, et al. Pyruvate kinase deficiency of mice associated with nonspherocytic hemolytic anemia and cure of the anemia by marrow transplantation without host irradiation. Blood. 1995;86(11):4323–30. pmid:7492793
- View Article
- PubMed/NCBI
- Google Scholar
51. WHO Organization. WHO Global Malaria Programme: World Malaria Report 2013. Geneva, Switzerland: WHO Press; 2013. p. 7
52. WHO Organization. Malaria: Fact sheet No94 Geneva, Switzerland: WHO Press; 2015 [cited 2015 October 2, 2015]. Available from: http://www.who.int/mediacentre/factsheets/fs094/en/.
53. Hedrick PW. Population genetics of malaria resistance in humans. Heredity (Edinb). 2011;107(4):283–304.
- View Article
- Google Scholar
54. Hedrick PW. Resistance to malaria in humans: the impact of strong, recent selection. Malar J. 2012;11:349. pmid:23088866
- View Article
- PubMed/NCBI
- Google Scholar
55. Smith T, Genton B, Baea K, Gibson N, Narara A, Alpers MP. Prospective risk of morbidity in relation to malaria infection in an area of high endemicity of multiple species of Plasmodium. Am J Trop Med Hyg. 2001;64(5–6):262–7. pmid:11463113
- View Article
- PubMed/NCBI
- Google Scholar
56. Zanella A, Fermo E, Bianchi P, Valentini G. Red cell pyruvate kinase deficiency: molecular and clinical aspects. Br J Haematol. 2005;130(1):11–25. pmid:15982340
- View Article
- PubMed/NCBI
- Google Scholar
57. Wang C, Chiarelli LR, Bianchi P, Abraham DJ, Galizzi A, Mattevi A, et al. Human erythrocyte pyruvate kinase: characterization of the recombinant enzyme and a mutant form (R510Q) causing nonspherocytic hemolytic anemia. Blood. 2001;98(10):3113–20. pmid:11698298
- View Article
- PubMed/NCBI
- Google Scholar
58. Kahn A, Marie J. Pyruvate kinases from human erythrocytes and liver. Methods Enzymol. 1982;90 Pt E:131–40. pmid:7154942
- View Article
- PubMed/NCBI
- Google Scholar
59. Marie J, Buc H, Simon MP, Kahn A. Phosphorylation of human erythrocyte pyruvate kinase by soluble cyclic-AMP-dependent protein kinases. Comparison with human liver L-type enzyme. Eur J Biochem. 1980;108(1):251–60. pmid:6250830
- View Article
- PubMed/NCBI
- Google Scholar
60. Fermo E, Bianchi P, Chiarelli LR, Cotton F, Vercellati C, Writzl K, et al. Red cell pyruvate kinase deficiency: 17 new mutations of the PK-LR gene. Br J Haematol. 2005;129(6):839–46. pmid:15953013
- View Article
- PubMed/NCBI
- Google Scholar
61. Zurcher C, Loos JA, Prins HK. Hereditary high ATP content of human erythrocytes. Folia Haematol Int Mag Klin Morphol Blutforsch. 1965;83(4):366–76. pmid:4160306
- View Article
- PubMed/NCBI
- Google Scholar
62. Beutler E, Westwood B, van Zwieten R, Roos D. G—>T transition at cDNA nt 110 (K37Q) in the PKLR (pyruvate kinase) gene is the molecular basis of a case of hereditary increase of red blood cell ATP. Hum Mutat. 1997;9(3):282–5. pmid:9090535
- View Article
- PubMed/NCBI
- Google Scholar
63. Marie J, Kahn A. Proteolytic processing of human erythrocyte pyruvate kinase: study of normal and deficient enzymes. Biochem Biophys Res Commun. 1979;91(1):123–9. pmid:518614
- View Article
- PubMed/NCBI
- Google Scholar
64. Fanjul AN, Farias RN. Cold-sensitive cytosolic 3,5,3'-triiodo-L-thyronine-binding protein and pyruvate kinase from human erythrocytes share similar regulatory properties of hormone binding by glycolytic intermediates. J Biol Chem. 1993;268(1):175–9. pmid:8416925
- View Article
- PubMed/NCBI
- Google Scholar
65. Gupta V, Bamezai RN. Human pyruvate kinase M2: a multifunctional protein. Protein Sci. 2010;19(11):2031–44. pmid:20857498
- View Article
- PubMed/NCBI
- Google Scholar
66. Strandholm JJ, Dyson RD, Cardenas JM. Bovine pyruvate kinase isozymes and hybrid isozymes. Electrophoretic studies and tissue distribution. Arch Biochem Biophys. 1976;173(1):125–31. pmid:1259434
- View Article
- PubMed/NCBI
- Google Scholar
67. Cardenas JM, Dyson RD. Mammalian pyruvate kinase hybrid isozymes: tissue distribution and physiological significance. J Exp Zool. 1978;204(3):361–7. pmid:660140
- View Article
- PubMed/NCBI
- Google Scholar
68. Cardenas JM, Richards TC, Gabourel L. Localization of pyruvate kinase isozymes in bovine kidney and comparison of these patterns with those of lactate dehydrogenases and aldolases. J Cell Physiol. 1978;96(2):189–97. pmid:670304
- View Article
- PubMed/NCBI
- Google Scholar
69. Mueggler PA, Carpenter S, Black JA. Postnatal changes in canine erythrocyte pyruvate kinase isozymes. Biochem Genet. 1983;21(7–8):691–702. pmid:6626141
- View Article
- PubMed/NCBI
- Google Scholar

[ref1] 1. Kwiatkowski DP. How malaria has affected the human genome and what human genetics can teach us about malaria. Am J Hum Genet. 2005;77(2):171–92. pmid:16001361
View Article
PubMed/NCBI
Google Scholar

[2] View Article

[3] PubMed/NCBI

[4] Google Scholar

[ref2] 2. Bongfen SE, Laroque A, Berghout J, Gros P. Genetic and genomic analyses of host-pathogen interactions in malaria. Trends Parasitol. 2009;25(9):417–22. pmid:19717339
View Article
PubMed/NCBI
Google Scholar

[6] View Article

[7] PubMed/NCBI

[8] Google Scholar

[ref3] 3. Min-Oo G, Gros P. Erythrocyte variants and the nature of their malaria protective effect. Cell Microbiol. 2005;7(6):753–63. pmid:15888079
View Article
PubMed/NCBI
Google Scholar

[10] View Article

[11] PubMed/NCBI

[12] Google Scholar

[ref4] 4. Williams TN, Wambua S, Uyoga S, Macharia A, Mwacharo JK, Newton CR, et al. Both heterozygous and homozygous alpha+ thalassemias protect against severe and fatal Plasmodium falciparum malaria on the coast of Kenya. Blood. 2005;106(1):368–71. pmid:15769889
View Article
PubMed/NCBI
Google Scholar

[14] View Article

[15] PubMed/NCBI

[16] Google Scholar

[ref5] 5. Rihet P, Traore Y, Abel L, Aucan C, Traore-Leroux T, Fumoux F. Malaria in humans: Plasmodium falciparum blood infection levels are linked to chromosome 5q31-q33. Am J Hum Genet. 1998;63(2):498–505. pmid:9683598
View Article
PubMed/NCBI
Google Scholar

[18] View Article

[19] PubMed/NCBI

[20] Google Scholar

[ref6] 6. Phimpraphi W, Paul R, Witoonpanich B, Turbpaiboon C, Peerapittayamongkol C, Louicharoen C, et al. Heritability of P. falciparum and P. vivax malaria in a Karen population in Thailand. PLoS One. 2008;3(12):e3887. pmid:19060954
View Article
PubMed/NCBI
Google Scholar

[22] View Article

[23] PubMed/NCBI

[24] Google Scholar

[ref7] 7. Lawaly YR, Sakuntabhai A, Marrama L, Konate L, Phimpraphi W, Sokhna C, et al. Heritability of the human infectious reservoir of malaria parasites. PLoS One. 2010;5(6):e11358. pmid:20613877
View Article
PubMed/NCBI
Google Scholar

[26] View Article

[27] PubMed/NCBI

[28] Google Scholar

[ref8] 8. Sakuntabhai A, Ndiaye R, Casademont I, Peerapittayamongkol C, Rogier C, Tortevoye P, et al. Genetic determination and linkage mapping of Plasmodium falciparum malaria related traits in Senegal. PLoS One. 2008;3(4):e2000. pmid:18431485
View Article
PubMed/NCBI
Google Scholar

[30] View Article

[31] PubMed/NCBI

[32] Google Scholar

[ref9] 9. Timmann C, Evans JA, Konig IR, Kleensang A, Ruschendorf F, Lenzen J, et al. Genome-wide linkage analysis of malaria infection intensity and mild disease. PLoS Genet. 2007;3(3):e48. pmid:17381244
View Article
PubMed/NCBI
Google Scholar

[34] View Article

[35] PubMed/NCBI

[36] Google Scholar

[ref10] 10. Jallow M, Teo YY, Small KS, Rockett KA, Deloukas P, Clark TG, et al. Genome-wide and fine-resolution association analysis of malaria in West Africa. Nat Genet. 2009;41(6):657–65. pmid:19465909
View Article
PubMed/NCBI
Google Scholar

[38] View Article

[39] PubMed/NCBI

[40] Google Scholar

[ref11] 11. Louicharoen C, Patin E, Paul R, Nuchprayoon I, Witoonpanich B, Peerapittayamongkol C, et al. Positively selected G6PD-Mahidol mutation reduces Plasmodium vivax density in Southeast Asians. Science. 2009;326(5959):1546–9. pmid:20007901
View Article
PubMed/NCBI
Google Scholar

[42] View Article

[43] PubMed/NCBI

[44] Google Scholar

[ref12] 12. Zimmerman PA, Woolley I, Masinde GL, Miller SM, McNamara DT, Hazlett F, et al. Emergence of FY*A(null) in a Plasmodium vivax-endemic region of Papua New Guinea. Proc Natl Acad Sci U S A. 1999;96(24):13973–7. pmid:10570183
View Article
PubMed/NCBI
Google Scholar

[46] View Article

[47] PubMed/NCBI

[48] Google Scholar

[ref13] 13. Miller LH, Mason SJ, Clyde DF, McGinniss MH. The resistance factor to Plasmodium vivax in blacks. The Duffy-blood-group genotype, FyFy. N Engl J Med. 1976;295(6):302–4. pmid:778616
View Article
PubMed/NCBI
Google Scholar

[50] View Article

[51] PubMed/NCBI

[52] Google Scholar

[ref14] 14. Maier AG, Duraisingh MT, Reeder JC, Patel SS, Kazura JW, Zimmerman PA, et al. Plasmodium falciparum erythrocyte invasion through glycophorin C and selection for Gerbich negativity in human populations. Nat Med. 2003;9(1):87–92. pmid:12469115
View Article
PubMed/NCBI
Google Scholar

[54] View Article

[55] PubMed/NCBI

[56] Google Scholar

[ref15] 15. Genton B, al-Yaman F, Mgone CS, Alexander N, Paniu MM, Alpers MP, et al. Ovalocytosis and cerebral malaria. Nature. 1995;378(6557):564–5. pmid:8524388
View Article
PubMed/NCBI
Google Scholar

[58] View Article

[59] PubMed/NCBI

[60] Google Scholar

[ref16] 16. Friedman MJ. Erythrocytic mechanism of sickle cell resistance to malaria. Proc Natl Acad Sci U S A. 1978;75(4):1994–7. pmid:347452
View Article
PubMed/NCBI
Google Scholar

[62] View Article

[63] PubMed/NCBI

[64] Google Scholar

[ref17] 17. Ayi K, Turrini F, Piga A, Arese P. Enhanced phagocytosis of ring-parasitized mutant erythrocytes: a common mechanism that may explain protection against falciparum malaria in sickle trait and beta-thalassemia trait. Blood. 2004;104(10):3364–71. pmid:15280204
View Article
PubMed/NCBI
Google Scholar

[66] View Article

[67] PubMed/NCBI

[68] Google Scholar

[ref18] 18. Modiano D, Luoni G, Sirima BS, Lanfrancotti A, Petrarca V, Cruciani F, et al. The lower susceptibility to Plasmodium falciparum malaria of Fulani of Burkina Faso (west Africa) is associated with low frequencies of classic malaria-resistance genes. Trans R Soc Trop Med Hyg. 2001;95(2):149–52. pmid:11355545
View Article
PubMed/NCBI
Google Scholar

[70] View Article

[71] PubMed/NCBI

[72] Google Scholar

[ref19] 19. Olivieri NF, Muraca GM, O'Donnell A, Premawardhena A, Fisher C, Weatherall DJ. Studies in haemoglobin E beta-thalassaemia. Br J Haematol. 2008;141(3):388–97. pmid:18410572
View Article
PubMed/NCBI
Google Scholar

[74] View Article

[75] PubMed/NCBI

[76] Google Scholar

[ref20] 20. Mockenhaupt FP, Ehrhardt S, Gellert S, Otchwemah RN, Dietz E, Anemana SD, et al. Alpha(+)-thalassemia protects African children from severe malaria. Blood. 2004;104(7):2003–6. pmid:15198952
View Article
PubMed/NCBI
Google Scholar

[78] View Article

[79] PubMed/NCBI

[80] Google Scholar

[ref21] 21. Rowe JA, Opi DH, Williams TN. Blood groups and malaria: fresh insights into pathogenesis and identification of targets for intervention. Curr Opin Hematol. 2009;16(6):480–7. pmid:19812491
View Article
PubMed/NCBI
Google Scholar

[82] View Article

[83] PubMed/NCBI

[84] Google Scholar

[ref22] 22. Fry AE, Griffiths MJ, Auburn S, Diakite M, Forton JT, Green A, et al. Common variation in the ABO glycosyltransferase is associated with susceptibility to severe Plasmodium falciparum malaria. Hum Mol Genet. 2008;17(4):567–76. pmid:18003641
View Article
PubMed/NCBI
Google Scholar

[86] View Article

[87] PubMed/NCBI

[88] Google Scholar

[ref23] 23. van Wijk R, van Solinge WW. The energy-less red blood cell is lost: erythrocyte enzyme abnormalities of glycolysis. Blood. 2005;106(13):4034–42. pmid:16051738
View Article
PubMed/NCBI
Google Scholar

[90] View Article

[91] PubMed/NCBI

[92] Google Scholar

[ref24] 24. Zanella A, Bianchi P. Red cell pyruvate kinase deficiency: from genetics to clinical manifestations. Baillieres Best Pract Res Clin Haematol. 2000;13(1):57–81. pmid:10916678
View Article
PubMed/NCBI
Google Scholar

[94] View Article

[95] PubMed/NCBI

[96] Google Scholar

[ref25] 25. Kanno H, Fujii H, Miwa S. Structural analysis of human pyruvate kinase L-gene and identification of the promoter activity in erythroid cells. Biochem Biophys Res Commun. 1992;188(2):516–23. pmid:1445295
View Article
PubMed/NCBI
Google Scholar

[98] View Article

[99] PubMed/NCBI

[100] Google Scholar

[ref26] 26. van Wijk R, Huizinga EG, van Wesel AC, van Oirschot BA, Hadders MA, van Solinge WW. Fifteen novel mutations in PKLR associated with pyruvate kinase (PK) deficiency: structural implications of amino acid substitutions in PK. Hum Mutat. 2009;30(3):446–53. pmid:19085939
View Article
PubMed/NCBI
Google Scholar

[102] View Article

[103] PubMed/NCBI

[104] Google Scholar

[ref27] 27. Gupta RK, Oesterling RM. Dual divalent cation requirement for activation of pyruvate kinase; essential roles of both enzyme- and nucleotide-bound metal ions. Biochemistry. 1976;15(13):2881–7. pmid:7293
View Article
PubMed/NCBI
Google Scholar

[106] View Article

[107] PubMed/NCBI

[108] Google Scholar

[ref28] 28. Valentine WN, Tanaka KR, Miwa S. A specific erythrocyte glycolytic enzyme defect (pyruvate kinase) in three subjects with congenital non-spherocytic hemolytic anemia. Trans Assoc Am Physicians. 1961;74:100–10. pmid:13924348
View Article
PubMed/NCBI
Google Scholar

[110] View Article

[111] PubMed/NCBI

[112] Google Scholar

[ref29] 29. Hirono A, Kanno H, Miwa S, Beutler E. Pyruvate kinase deficiency and other enzymopathies of the erythrocyte. In: Scriver CR, editor. The Metabolic and Molecular Bases of Inherited Disease. 8th ed ed. New York: McGraw-Hill; 2001. p. 4637–64.

[ref30] 30. Ayi K, Min-Oo G, Serghides L, Crockett M, Kirby-Allen M, Quirt I, et al. Pyruvate kinase deficiency and malaria. N Engl J Med. 2008;358(17):1805–10. pmid:18420493
View Article
PubMed/NCBI
Google Scholar

[115] View Article

[116] PubMed/NCBI

[117] Google Scholar

[ref31] 31. Min-Oo G, Fortin A, Tam MF, Gros P, Stevenson MM. Phenotypic expression of pyruvate kinase deficiency and protection against malaria in a mouse model. Genes Immun. 2004;5(3):168–75. pmid:15029238
View Article
PubMed/NCBI
Google Scholar

[119] View Article

[120] PubMed/NCBI

[121] Google Scholar

[ref32] 32. Min-Oo G, Fortin A, Tam MF, Nantel A, Stevenson MM, Gros P. Pyruvate kinase deficiency in mice protects against malaria. Nat Genet. 2003;35(4):357–62. pmid:14595440
View Article
PubMed/NCBI
Google Scholar

[123] View Article

[124] PubMed/NCBI

[125] Google Scholar

[ref33] 33. Durand PM, Coetzer TL. Pyruvate kinase deficiency protects against malaria in humans. Haematologica. 2008;93(6):939–40. pmid:18460648
View Article
PubMed/NCBI
Google Scholar

[127] View Article

[128] PubMed/NCBI

[129] Google Scholar

[ref34] 34. Berghout J, Higgins S, Loucoubar C, Sakuntabhai A, Kain KC, Gros P. Genetic diversity in human erythrocyte pyruvate kinase. Genes Immun. 2012;13(1):98–102. pmid:21833022
View Article
PubMed/NCBI
Google Scholar

[131] View Article

[132] PubMed/NCBI

[133] Google Scholar

[ref35] 35. Machado P, Manco L, Gomes C, Mendes C, Fernandes N, Salome G, et al. Pyruvate kinase deficiency in sub-Saharan Africa: identification of a highly frequent missense mutation (G829A;Glu277Lys) and association with malaria. PLoS One. 2012;7(10):e47071. pmid:23082140
View Article
PubMed/NCBI
Google Scholar

[135] View Article

[136] PubMed/NCBI

[137] Google Scholar

[ref36] 36. Alves J, Machado P, Silva J, Goncalves N, Ribeiro L, Faustino P, et al. Analysis of malaria associated genetic traits in Cabo Verde, a melting pot of European and sub Saharan settlers. Blood Cells Mol Dis. 2010;44(1):62–8. pmid:19837619
View Article
PubMed/NCBI
Google Scholar

[139] View Article

[140] PubMed/NCBI

[141] Google Scholar

[ref37] 37. Phimpraphi W, Paul RE, Yimsamran S, Puangsa-art S, Thanyavanich N, Maneeboonyang W, et al. Longitudinal study of Plasmodium falciparum and Plasmodium vivax in a Karen population in Thailand. Malar J. 2008;7:99. pmid:18518964
View Article
PubMed/NCBI
Google Scholar

[143] View Article

[144] PubMed/NCBI

[145] Google Scholar

[ref38] 38. Trape JF, Rogier C, Konate L, Diagne N, Bouganali H, Canque B, et al. The Dielmo project: a longitudinal study of natural malaria infection and the mechanisms of protective immunity in a community living in a holoendemic area of Senegal. Am J Trop Med Hyg. 1994;51(2):123–37. pmid:8074247
View Article
PubMed/NCBI
Google Scholar

[147] View Article

[148] PubMed/NCBI

[149] Google Scholar

[ref39] 39. Rogier C, Commenges D, Trape JF. Evidence for an age-dependent pyrogenic threshold of Plasmodium falciparum parasitemia in highly endemic populations. Am J Trop Med Hyg. 1996;54(6):613–9. pmid:8686780
View Article
PubMed/NCBI
Google Scholar

[151] View Article

[152] PubMed/NCBI

[153] Google Scholar

[ref40] 40. Rogier C, Ly AB, Tall A, Cisse B, Trape JF. Plasmodium falciparum clinical malaria in Dielmo, a holoendemic area in Senegal: no influence of acquired immunity on initial symptomatology and severity of malaria attacks. Am J Trop Med Hyg. 1999;60(3):410–20. pmid:10466970
View Article
PubMed/NCBI
Google Scholar

[155] View Article

[156] PubMed/NCBI

[157] Google Scholar

[ref41] 41. Rogier C, Tall A, Diagne N, Fontenille D, Spiegel A, Trape JF. Plasmodium falciparum clinical malaria: lessons from longitudinal studies in Senegal. Parassitologia. 1999;41(1–3):255–9. pmid:10697865
View Article
PubMed/NCBI
Google Scholar

[159] View Article

[160] PubMed/NCBI

[161] Google Scholar

[ref42] 42. Hall TA. BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symposium Series. 1999;41:95–8.
View Article
Google Scholar

[163] View Article

[164] Google Scholar

[ref43] 43. Barrett JC, Fry B, Maller J, Daly MJ. Haploview: analysis and visualization of LD and haplotype maps. Bioinformatics. 2005;21(2):263–5. pmid:15297300
View Article
PubMed/NCBI
Google Scholar

[166] View Article

[167] PubMed/NCBI

[168] Google Scholar

[ref44] 44. O'Connell JR, Weeks DE. PedCheck: a program for identification of genotype incompatibilities in linkage analysis. Am J Hum Genet. 1998;63(1):259–66. pmid:9634505
View Article
PubMed/NCBI
Google Scholar

[170] View Article

[171] PubMed/NCBI

[172] Google Scholar

[ref45] 45. Fortin A, Cardon LR, Tam M, Skamene E, Stevenson MM, Gros P. Identification of a new malaria susceptibility locus (Char4) in recombinant congenic strains of mice. Proc Natl Acad Sci U S A. 2001;98(19):10793–8. pmid:11535821
View Article
PubMed/NCBI
Google Scholar

[174] View Article

[175] PubMed/NCBI

[176] Google Scholar

[ref46] 46. Valentini G, Chiarelli LR, Fortin R, Dolzan M, Galizzi A, Abraham DJ, et al. Structure and function of human erythrocyte pyruvate kinase. Molecular basis of nonspherocytic hemolytic anemia. J Biol Chem. 2002;277(26):23807–14. pmid:11960989
View Article
PubMed/NCBI
Google Scholar

[178] View Article

[179] PubMed/NCBI

[180] Google Scholar

[ref47] 47. Modiano D, Petrarca V, Sirima BS, Bosman A, Nebie I, Diallo D, et al. Plasmodium falciparum malaria in sympatric ethnic groups of Burkina Faso, west Africa. Parassitologia. 1995;37(2–3):255–9. pmid:8778668
View Article
PubMed/NCBI
Google Scholar

[182] View Article

[183] PubMed/NCBI

[184] Google Scholar

[ref48] 48. Tiono AB, Sirima SB, Hamed K. Fulani show decreased susceptibility to Plasmodium falciparum infection versus Mossi: data from a community-wide screening and treatment of asymptomatic carriers in Burkina Faso. Malar J. 2013;12:163. pmid:23680454
View Article
PubMed/NCBI
Google Scholar

[186] View Article

[187] PubMed/NCBI

[188] Google Scholar

[ref49] 49. Summerer M, Horst J, Erhart G, Weissensteiner H, Schonherr S, Pacher D, et al. Large-scale mitochondrial DNA analysis in Southeast Asia reveals evolutionary effects of cultural isolation in the multi-ethnic population of Myanmar. BMC Evol Biol. 2014;14:17. pmid:24467713
View Article
PubMed/NCBI
Google Scholar

[190] View Article

[191] PubMed/NCBI

[192] Google Scholar

[ref50] 50. Morimoto M, Kanno H, Asai H, Tsujimura T, Fujii H, Moriyama Y, et al. Pyruvate kinase deficiency of mice associated with nonspherocytic hemolytic anemia and cure of the anemia by marrow transplantation without host irradiation. Blood. 1995;86(11):4323–30. pmid:7492793
View Article
PubMed/NCBI
Google Scholar

[194] View Article

[195] PubMed/NCBI

[196] Google Scholar

[ref51] 51. WHO Organization. WHO Global Malaria Programme: World Malaria Report 2013. Geneva, Switzerland: WHO Press; 2013. p. 7

[ref52] 52. WHO Organization. Malaria: Fact sheet No94 Geneva, Switzerland: WHO Press; 2015 [cited 2015 October 2, 2015]. Available from: http://www.who.int/mediacentre/factsheets/fs094/en/.

[ref53] 53. Hedrick PW. Population genetics of malaria resistance in humans. Heredity (Edinb). 2011;107(4):283–304.
View Article
Google Scholar

[200] View Article

[201] Google Scholar

[ref54] 54. Hedrick PW. Resistance to malaria in humans: the impact of strong, recent selection. Malar J. 2012;11:349. pmid:23088866
View Article
PubMed/NCBI
Google Scholar

[203] View Article

[204] PubMed/NCBI

[205] Google Scholar

[ref55] 55. Smith T, Genton B, Baea K, Gibson N, Narara A, Alpers MP. Prospective risk of morbidity in relation to malaria infection in an area of high endemicity of multiple species of Plasmodium. Am J Trop Med Hyg. 2001;64(5–6):262–7. pmid:11463113
View Article
PubMed/NCBI
Google Scholar

[207] View Article

[208] PubMed/NCBI

[209] Google Scholar

[ref56] 56. Zanella A, Fermo E, Bianchi P, Valentini G. Red cell pyruvate kinase deficiency: molecular and clinical aspects. Br J Haematol. 2005;130(1):11–25. pmid:15982340
View Article
PubMed/NCBI
Google Scholar

[211] View Article

[212] PubMed/NCBI

[213] Google Scholar

[ref57] 57. Wang C, Chiarelli LR, Bianchi P, Abraham DJ, Galizzi A, Mattevi A, et al. Human erythrocyte pyruvate kinase: characterization of the recombinant enzyme and a mutant form (R510Q) causing nonspherocytic hemolytic anemia. Blood. 2001;98(10):3113–20. pmid:11698298
View Article
PubMed/NCBI
Google Scholar

[215] View Article

[216] PubMed/NCBI

[217] Google Scholar

[ref58] 58. Kahn A, Marie J. Pyruvate kinases from human erythrocytes and liver. Methods Enzymol. 1982;90 Pt E:131–40. pmid:7154942
View Article
PubMed/NCBI
Google Scholar

[219] View Article

[220] PubMed/NCBI

[221] Google Scholar

[ref59] 59. Marie J, Buc H, Simon MP, Kahn A. Phosphorylation of human erythrocyte pyruvate kinase by soluble cyclic-AMP-dependent protein kinases. Comparison with human liver L-type enzyme. Eur J Biochem. 1980;108(1):251–60. pmid:6250830
View Article
PubMed/NCBI
Google Scholar

[223] View Article

[224] PubMed/NCBI

[225] Google Scholar

[ref60] 60. Fermo E, Bianchi P, Chiarelli LR, Cotton F, Vercellati C, Writzl K, et al. Red cell pyruvate kinase deficiency: 17 new mutations of the PK-LR gene. Br J Haematol. 2005;129(6):839–46. pmid:15953013
View Article
PubMed/NCBI
Google Scholar

[227] View Article

[228] PubMed/NCBI

[229] Google Scholar

[ref61] 61. Zurcher C, Loos JA, Prins HK. Hereditary high ATP content of human erythrocytes. Folia Haematol Int Mag Klin Morphol Blutforsch. 1965;83(4):366–76. pmid:4160306
View Article
PubMed/NCBI
Google Scholar

[231] View Article

[232] PubMed/NCBI

[233] Google Scholar

[ref62] 62. Beutler E, Westwood B, van Zwieten R, Roos D. G—>T transition at cDNA nt 110 (K37Q) in the PKLR (pyruvate kinase) gene is the molecular basis of a case of hereditary increase of red blood cell ATP. Hum Mutat. 1997;9(3):282–5. pmid:9090535
View Article
PubMed/NCBI
Google Scholar

[235] View Article

[236] PubMed/NCBI

[237] Google Scholar

[ref63] 63. Marie J, Kahn A. Proteolytic processing of human erythrocyte pyruvate kinase: study of normal and deficient enzymes. Biochem Biophys Res Commun. 1979;91(1):123–9. pmid:518614
View Article
PubMed/NCBI
Google Scholar

[239] View Article

[240] PubMed/NCBI

[241] Google Scholar

[ref64] 64. Fanjul AN, Farias RN. Cold-sensitive cytosolic 3,5,3'-triiodo-L-thyronine-binding protein and pyruvate kinase from human erythrocytes share similar regulatory properties of hormone binding by glycolytic intermediates. J Biol Chem. 1993;268(1):175–9. pmid:8416925
View Article
PubMed/NCBI
Google Scholar

[243] View Article

[244] PubMed/NCBI

[245] Google Scholar

[ref65] 65. Gupta V, Bamezai RN. Human pyruvate kinase M2: a multifunctional protein. Protein Sci. 2010;19(11):2031–44. pmid:20857498
View Article
PubMed/NCBI
Google Scholar

[247] View Article

[248] PubMed/NCBI

[249] Google Scholar

[ref66] 66. Strandholm JJ, Dyson RD, Cardenas JM. Bovine pyruvate kinase isozymes and hybrid isozymes. Electrophoretic studies and tissue distribution. Arch Biochem Biophys. 1976;173(1):125–31. pmid:1259434
View Article
PubMed/NCBI
Google Scholar

[251] View Article

[252] PubMed/NCBI

[253] Google Scholar

[ref67] 67. Cardenas JM, Dyson RD. Mammalian pyruvate kinase hybrid isozymes: tissue distribution and physiological significance. J Exp Zool. 1978;204(3):361–7. pmid:660140
View Article
PubMed/NCBI
Google Scholar

[255] View Article

[256] PubMed/NCBI

[257] Google Scholar

[ref68] 68. Cardenas JM, Richards TC, Gabourel L. Localization of pyruvate kinase isozymes in bovine kidney and comparison of these patterns with those of lactate dehydrogenases and aldolases. J Cell Physiol. 1978;96(2):189–97. pmid:670304
View Article
PubMed/NCBI
Google Scholar

[259] View Article

[260] PubMed/NCBI

[261] Google Scholar

[ref69] 69. Mueggler PA, Carpenter S, Black JA. Postnatal changes in canine erythrocyte pyruvate kinase isozymes. Biochem Genet. 1983;21(7–8):691–702. pmid:6626141
View Article
PubMed/NCBI
Google Scholar

[263] View Article

[264] PubMed/NCBI

[265] Google Scholar

Figures

Abstract

Introduction

Materials and Methods

Population Studies

Association studies and statistical analysis

Preparation of wild-type and mutant PKLR constructs

Protein Expression Studies

Protein Stability Assay

Mice and parasite

Results

PKLR genotypes in informative Senegalese and Thai populations

Population genetic analysis of the R41Q and L272V variants in PKLR

Association studies

Heterozygosity for Pklr in mice protects against peak parasitemia

R41Q recombinant protein exhibits modest effect on half-life

Discussion

Supporting Information

S1 Table. Sequencing primers designed to span the exon and intron/exon junctions of the PKLR gene.

S2 Table. Main characteristic of the Thai cohort (n = 897) investigated in this study.

S3 Table. List of haematological and malaria phenotypes investigated in this study.

S4 Table. Site directed mutagenesis primers.

S5 Table. Construct sequencing primers.

Acknowledgments

Author Contributions

References