Degree project 15 hp
Biomedical science programme, 180 hp
Department of Medical Biochemistry and Microbiology,
Uppsala University
Variation at position 86 of the pfmdr1 gene in samples from
an area with seasonal transmission in eastern Sudan
Tamara Villalta Montoya
Supervisors: Göte Swedberg and Amani Kheir,
Department of Medical Biochemistry and Microbiology,
Uppsala University
ABSTRACT
Malaria is the most common parasitic disease of humans worldwide. A factor that aggravates the many attempts to control the epidemiologic malaria situation is the spreading of resistance against anti-malarial drugs. In this project the point mutation at position 86 of the Plasmodium. falciparum multidrug resistance gene (pfmdr1), which is thought to contribute to Chloroquine resistance, was analysed in 188 samples from a low transmission area in eastern Sudan, where malaria endemicity is seasonal. The patient group studied had asymptomatic and sub patent parasitemia that persisted during the transmission-free dry season, after being treated with Chloroquine. To differentiate between wild type and mutant genotypes, nested PCR and restriction fragment length
polymorphism with the enzyme Apo1 was used. Out of 188 samples 79 (42%) were successfully analysed. Of those, 72% had parasites with mutant genotypes or where mixed infection. No conclusions on the relevance of the pfmdr1 gene in the studied samples are made due to the many remaining gaps. However, eventual sources of error and previous findings in the study area are discussed.
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INTRODUCTIONAfter being eliminated from a big part of the world, malaria is still one of the world’s biggest challenges, as it undoubtedly is the most common parasitic infection of humans worldwide. This makes it a big human and economic burden due to lack of control and high mortality rates in endemic areas. The World Health Organization (WHO) estimates that half of the world’s population was at risk of getting malaria in 2006, and the disease caused nearly a million deaths the same year, of which 85% were children under 5 years of age. Africa had by far the heaviest death incidence with as much as 91% of all the deaths (WHO malaria report 2008).
Malaria is an ancient disease, mentioned in several old scripts. The disease is caused by different protozoans of the genus Plasmodium, mainly Plasmodium falciparum, and is transmitted by the bite of infected female
Anopheles mosquitoes. The parasites have
different complex lifecycles in both human and in its vector. In humans the parasites enters the body as sporozoites, then travel through the bloodstream to invade hepatic cells, where they have an asexual reproduction stage, developing into merozoites. When the hepatocyte bursts, the merozoites are released into the bloodstream where they infect red blood cells (RBC). In this stage another asexual reproduction cycle takes place. Due to the asexual reproduction stages, parasites are haploid during the human lifecycle. The human cycle eventually ends when some of the merozoites develop into gametocytes, the form that can be transmitted to mosquitoes. The effects and consequences of the RBCs invasion, together with the immune system response, directly cause the medical conditions of malaria. The parasites can for instance induce surface modifications of the RBCs making them produce proteins that can adhere to the blood
vessel lining, a mechanism that enables the parasites to escape destruction in the spleen. But it also leads to thrombosis and local ischemia when the RBCs adhere to each other.
Clinical malaria episodes with P.falciparum, start after an incubation time of 2-4 weeks, with nonspecific symptoms similar to those of minor viral illness, such as fever and malaise, followed by symptoms like mild anemia and sometimes splenomegaly. Untreated, uncomplicated malaria, when the patient can swallow medicines and food, can develop into complicated malaria, also called severe malaria. If so, the mortality risk increases remarkably. Clinical features of complicated malaria involve vital organ dysfunction such as cerebral malaria, severe anemia, acidosis, renal impairment and liver dysfunction.
Malaria is mainly diagnosed through clinical signs like fever, fever history, palpitation of the spleen and pallor of the hands in children. People that live in endemic areas often know the clinical signs of malaria and consequently often diagnose themselves. In areas with laboratory facilities, the malaria diagnosis is supplemented by microscopic study of blood smears, or by rapid diagnostic tests (WHO Guidelines for the treatment of malaria).
therefore lost when the person moves from the area.
When treating patients with uncomplicated malaria the main goal is to eliminate the parasites from the body and hence cure the infection to prevent a progression to severe
malaria. When treating severe malaria the main objective is to prevent death. Today Artemisinin based combination therapy is recommended as the first line defence. The emergence of drug resistance has lead to the conclusion that monotherapy is no longer recommended (WHO Guidelines for the treatment of malaria). Other drugs that have been, and still are used, are Chloroquine (CQ) and related compounds, antifolates such as sulfadoxine-pyrimethamine (SP), and Atovaquone (Hyde 2007).
After using CQ as the drug of choice for nearly 20 years, resistance arose, and reports started to come from different parts of the world during the 1960s. Today the resistance to CQ is so widely spread, that it is almost exactly equal to that of the parasite (Talisuna et al., 2004). Two genes are related to the resistance and are believed to be responsible for the mechanisms. One is the P.
falciparum CQ resistance related transporter
protein gene (pfcrt) situated in chromosome 7 (Fidock et al., 2000), and the other is the P.
falciparum multidrug resistance gene (pfmdr1) on
chromosome 5; a gene that encodes for P-glycoprotein 1 (Foote et al., 1990). Much evidence suggest that the mutations in the pfcrt gene plays the biggest role in resistance to CQ, especially a point mutation in position 76 that gives a substitution of threonine to lysine (Talisuna et al., 2004; Hyde 2007). Although the role of the pfmdr1 gene has been widely studied around the world, the conclusions are not consistent. Some studies strengthen the association between pfmdr1 and CQ resistance (Duraisingh et al., 1996, Foote et al., 1990), while others do not find any evidence supporting the
linkage (Pillai et al., 2001; Thomas et al., 2002). In addition to those studies there are further studies where the conclusion remains unclear (Basco et al., 1995; Djimdé et al., 2001). The general conclusion today is that the mutations of the pfmdr1 gene leads to CQ resistance, but that the resulting amino acid changes are not solely responsible for the resistance to the drug and clinical failure (Babiker et al., 2001; Hyde 2007; Talisuna et al., 2004). Five different point mutations within the pfmrd1 gene are associated with CQ resistance (Foote et al., 1990). The mutation that is mostly associated with the resistance is a mutation in position 86 that results in an amino acid change form aspargine to tyrosine (Duraisingh et al., 1996; Talisuna et al., 2004). Other mutations that are linked to the resistance are found in position 184, 1034, 1042 and 1246. These mutations also give rise to amino acid changes (Foote et al., 1990). When it comes to the biochemical mechanism of CQ resistance, it is thought to work by lowering the drug accumulation in the parasites digestive organ, the equivalent of the human lysosome (Duraisingh et al., 2005).
This project was a continuation of previous work done on samples collected from inhabitants from the village of Asar, in the eastern part of Sudan. This is an area where several studies have been done during different time periods (Babiker et al., 2005; Abdel-Muhsin et al., 2004). Asar is a low transmission area where the malaria transmission is seasonal, taking place during August to November each year after the rain season, when there is a brief expansion of the
Anopheles arabiensis mosquito, which is the
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are at risk of malaria in low transmission areas, due to a setback of the partial immunity (WHO Guidelines for the treatment of malaria). The annual entomological inoculation rate (EIR); i.e. how many times a person is bitten by an infected mosquito, is around two to three during the transmission period, and thereby low (Hamad et al., 2002).
The project is based on prior findings on samples collected during the time period of October 1993 to December 1994. Out of a larger patient group, in which all had uncomplicated malaria, and were treated with QC or SP, three cohort patient groups were identified. The first group cleared their infections after treatment. The second group also cleared their infections, but was reinfected in the following transmission season. The third group did not clear their infections at all. These patients went from having symptomatic and patent infections, to having chronic and asymptomatic infections that ranged from patent to subpatent (Babiker et al., 1998). A patent infection is when the parasitemia is detectable through studying blood smears by light microscope, and a subpatent infection is when no parasites are found in the blood smear, but by another assay, such as polymerase chain reaction (PCR).
Almost all of the patients in the third group had initially mixed infections (i.e. infections caused by coexisting parasites) that during the whole study period varied in combination. All of them had some, or all, of the initial clones when reinvestigated in 1994, and some of the patients also showed infections with new parasites. Some of the patients had clinical episodes following the first episode, although almost all of the patients where symptom free during the period of March and September 1994 (Babiker et al., 1998). The purpose of this project was to study if the parasites in the samples from these patients had the point mutation at position 86 of the pfmdr1
gene that is associated with CQ resistance, to answer the question whether resistance to QC is part of the explanation to why parasites survived the transmission and drug-free season of 1994 or not. It is part of a larger, ongoing project, where the objective is to investigate the characteristics and underlying factors to the evolution of drug resistant strains in eastern Sudan, making this project a piece of a big puzzle.
MATERIALS AND METHODS Samples
The 188 samples that were included in this project came from 22 habitants of the village of Asar in eastern Sudan. The samples were collected during a time period of 15 months, from October 1993 to December 1994. Ethical clearance was obtained from The Ethical Clearance Committee of the Ministry of Health of Sudan and informal consent was obtained. All patients were first diagnosed with P. falciparum malaria by microscopy of blood smears, and then treated with a standard dose of CQ (25 mg/kg). Subsequently, finger pricked blood samples were regularly collected during the following 15 months, that is during the transmission season of 1993, the dry season of 1994 and the transmission season of 1994. During October 1993 two samples were taken. The samples were centrifuged and stored in liquid nitrogen (Babiker et al., 1998). In 2006 the samples were spotted on Whatman # 3 filter papers (Whatman plc) to facilitate their transport.
DNA extraction
Reference strains
The CQ sensitive 3D7 and the CQ resistant Dd2 were cultivated in vitro at the University of Edinburgh.
Amplification of position 86 with Nested PCR
The PCR assay used for amplification of the target position was a nested PCR. It consisted of two different complementary reactions, one outer reaction, and one nested reaction, where the PCR product from the outer reaction was used as template in the nested reaction. The reactions were carried out in a Mastercycler gradient from Eppendorf.
This assay has been used for malaria detection from field samples collected on filter paper since the middle of the 90s. It has a high specificity and sensitivity compared to standard microscopy, making it possible to detect very low parasite amount (Singh et al., 1996).
Procedure
The basic temperature cycles were the same for both reactions, with an initializing step for 3 min at 94 C, denaturation for 1 min at 94 C, annealing for 1 min at 45 C, elongation for 1 min at 72 C, final elongation for 10 min at 72 C and a final hold at 4 C. Only the number of cycles differed. The outer reaction needed 34 cycles and the nested reaction needed 30 cycles.
To evaluate the PCR, the product was detected using a 1.5% agarose gel stained with 5-7% ethidium bromide. It was run for 30 min at 90V. The result was then compared with a 100 – 1000 bp ladder (Fermentas) to estimate the size of the product (559 bp).
Reagents used for the Amplification
There were a number of difficulties connected to the amplifications. Therefore, two different DNA polymerase kits and three different protocols were used.
a) For the majority of samples (n=97) the DreamTaq™ Green DNA Polymerase kit (Fermentas) was used with a 2.5 Unit DNA polymerase protocol. Each 20 µL reaction had a concentration of 1 X DreamTaq buffer (Fermentas), 200 µM dNTP mix (Roche Diagnostics), 0.2 pmol/µL from each primer solution, and 5 Units of DreamTaq DNA polymerase (Fermentas).
b) In a second group of samples (n=72) the DreamTaq™ Green DNA Polymerase kit was also used, but with a 5 Unit DNA polymerase protocol. Besides the amount of DNA polymerase, the protocol differed in dNTP concentration. Here the dNTP concentration was decreased to 100 µM. Also in this protocol a total reaction volume of 20 µL was used.
c) In the third group of samples (n = 19) the Illustra™ puReTaq Ready – To- Go PCR beads kit (GE healthcare) was used. Since the kit contained PCR tubes with beads having everything needed for the PCR except for sterile water and primers, these were added. As the manufactures recommended, a total volume of 25 µL was used per reaction. Each reaction had a final concentration of 10 mM Tris–HCl, pH 9.0 at room temperature, 50 mM KCl, 1.5 mM MgCl2,
200µM of each dNTP, 0.1 pmol/µL of each primer, and 2.5 Units of the puReTaq DNA polymerase
The volume of extracted plasmodial DNA used for all the reactions was: 2 µL for the outer PCR and 1µL for the nested PCR.
Primers
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(forward) and 5´ AAA GAT GGT AAC CTC AGT ATC AAA GAA GAG 3´ (reverse) (Babiker et al., 2001).
Restriction fragment length polymorphism
To differentiate between the wild type and the mutant genotype at position 86, restriction fragment length polymorphism with the enzyme Apo1 (New England Biolabs) was used. The wild type genotype was cut by the restriction enzyme at the designated restriction site resulting in two smaller fragments (254 bp, 235 bp), that were so close in size that they appeared as one band in the 1,5% agarose gel, when run at 90V for 30 min. The mutant genotype (TAT) was also cut at a site common to both the wild type and the mutant genotype, giving rise to a fragment of 78 bp. This was used as a control site to make sure that the enzyme was working (Duraisingh et al., 1996). Two different protocols were used for the digestion. When digesting samples amplified with the DreamTaq™ Green DNA Polymerase kit, a concentration of 1,5 X NEbuffer 3 (New England Biolabs), 1,5 X BSA (New England Biolabs) – both supplemented with the enzyme kit - and 2 Units of Apo1 was used. To digest the PCR product amplified with the Illustra™ puReTaq Ready – To- Go PCR beads kit, the manufactures instructions were as follows; 1 X BSA, 1 X NEbuffer 3 and 2 Units of Apo 1.
RESULTS
Out of 188 samples 79 (42%) were successfully amplified and digested. Of these, 64 samples were amplified with the DreamTaq™ Green
DNA Polymerase kit, of which 49 with the 5 Unit DNA polymerase protocol and 15 with the 2.5 Unit protocol. The remaining 15 samples were amplified using the Illustra™ puReTaq Ready – To- Go PCR beads kit. In 52 of the analysed samples (66%), the parasites had a mutant genotype, and in 22 of the samples (28%) a wild type genotype. The remaining five samples (6%) showed parasites of both wild type and mutant type indicating mixed infections, in this case with both the wild type and mutant genotype (Figure1).
Figure 1. Variation at position 86 of the pfmdr 1 gene in 79 succesfully amplified samples.
66% 28%
6%
Table 1. Variation at positions 86 of the pfmdr1 gene within each patient from successfully amplified samples collected from the transmission season 1993 to the transmission season of 1994.
P at ie n t ID O ct . 9 3 O ct . 9 3 N ov .9 3 D ec . 9 3 Ja n . 9 4 F eb . 9 4 M ar . 9 4 A p r. 9 4 M ay . 9 4 Ju n . 9 4 Ju l. 94 A u g. 9 4 S ep t. 9 4 O ct . 9 4 N ov . 9 4 D ec . 9 4 A MIX M B M M M M M C M M M M M D M M E M M M M M M F M M M G M H M W W M W M I M M M W MIX J M M M M W W K MIX W W W W MIX L M M M M M M M N M O M W W W W P MIX M M M M M M Q R W M M S W M W W T M M W U V W W W Transmission
During the dry and drug free season, the majority of patients had persistent parasites with mutant genotypes (Table 1). The exceptions were six patients (H, I, K, O, S, T) that demonstrated wild type parasites during this period. Ten of the patients (A, H, I, J, K, O, P, R, S, T) had different parasite types that varied during the whole study period, and that fluctuated in genotype from mutant to wild type or mixed, and vice versa. Some of the patients showed remarkable patterns. Patient H had for instance mutant parasites during the transmission season, then wild type parasites during the dry season, which then went from wild type to mutant genotype during a month, and back to wild type again. Patient S also showed a similar pattern.
DISCUSSION
The initial aim of this project was to investigate all point mutations of the pfmdr1 gene. However, this initial objective could not be reached due to many obstacles. Of all the 188 samples, only 79 were successfully amplified covering position 86. When it came to amplifying the area covering the rest of the point mutations, none of the samples were successfully amplified. The reasons can be discussed, considering that the samples were amplified with primers targeting the merozoite surface protein 1 (MSP-1) and 2 (MSP-2), and glutamate rich protein (GLURP), in a previously study (Babiker et al., 1998). Those genes are commonly targeted by PCR to characterize different parasite populations (mixed infections), since they demonstrate population diversity in both length and sequence (Färnert et al., 2001). Thus, the question is why only 42 % of the samples were successfully analysed when targeted for the pfmdr1 gene?
The low yield of PCR product might be explained by poor quality DNA, giving the PCR a low sensitivity. Many factors in the handling of samples before DNA extraction could have
The failure to amplify many of the samples could also be explained if the blood spotted on the filter papers, or the filter paper piece cut for the DNA extraction, did not hold any parasites. The lower the parasitemia is, the bigger is the risk of not getting infected RBCs in the fraction of the sample chosen to extract, thus, the sample is not representative.
A lot of trouble- shooting was done to amplify the DNA targets in the samples, besides using different DNA polymerase kits and Units of DNA polymerase. Among them was to try different MgCl2 concentrations, to increase and decrease
the amount of template, and to use gradient PCR to optimise the annealing temperature. Another extraction method was also used (Bereczky et al., 2005). However, more troubleshooting can be done, that eventually results in a working PCR protocol.
One major problem during troubleshooting was that it was not possible to measure the concentration of plasmodial DNA in the samples. When extracting the DNA, both human and plasmodial DNA got extracted since the parasites were inside the RBCs. The extracted DNA consisted therefore of both host DNA and pathogen DNA, and could not be separated. It was thus not possible to know anything about the plasmodial DNA concentration in the samples, or even if there were any plasmodial DNA in the samples. One way of solving the problem would be to use primers to amplify a multi copy gene, since the sensitivity of the PCR also depends on the number of copies of the target gene. The more copies of the target gene, the bigger the total yield is. The small subunit ribosomal gene has been widely used as a target to detect malaria parasites since it is present as 4-8 copies in each parasite, and has conserved regions (Berry et al., 2005). Since it is easier to get a detectable yield amplifying a multi copy gene, this would probably give good information about the plasmodial DNA amount in the samples.
Regarding the choice of PCR assay, a nested PCR was the appropriate assay. Especially since the samples studied had very low parasite amount. The two PCR rounds yield more PCR product, thereby increasing the sensitivity of the assay. However, there are two drawbacks. Firstly, there is a high contamination risk when handling PCR product as template in the nested reaction. Secondly, there are some methodological disadvantages in detecting mixed infections with PCR in general when not using multiplex PCR or targeting specific genes for the purpose. The chances of detecting different coexisting parasite populations decrease due to the exponential nature of the PCR (i.e. that the amount of PCR product doubles after each cycle). There is a risk that a less dominating population does not get detected in a mixed infection, since the outcome will depend on the amplification of the template in the first few cycles. This competition effect makes the detection of mixed infections more difficult, giving “false negative” results (Arnot D. 1998). For that reason it could not be excluded that more of the samples were mixed infection, implying another pattern to the results.
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that has biggest influence on the development to CQ resistance, and that the pfmdr1 gene has an contributing role (Hyde 2007). A later study showed that there was a significant increase in parasites with mutant genotypes in the above-mentioned positions during a twelve-year period. Samples that were analysed in that study came from a period that stretched from the wet season of 1990 to the wet season of 2001, including the wet season samples from 1993 that were studied in this project (Abdel-Muhsin et al., 2004). These previous results coincide with the results obtained in this project, where the majority of samples had parasites with mutant genotypes for position 86 of the pfmdr1 gene (Figure 1). Totally 72% of the samples showed parasites with mutant genotypes, including the samples with mixed infections. The longitudinal study also showed a wet to dry season fluctuation of the pfcrt gene during the seasons of 1998 and 1999 (Abdel-Muhsin et al., 2004). It would be interesting to see if the samples studied in this project also showed a similar pattern.
All the samples from the patient group studied in this project had previously shown low parasitemia that persisted during the drug and transmission free season of 1994. The infections were complex, containing coexisting parasites that fluctuated in composition during the study period (Babiker et al., 1998). Hence, these previous observations explain the fluctuating patters that some of the patients exhibited, when shifting between parasites with mutant genotypes to parasites wild type genotypes, and sometimes back again (Table1). However, the methodological problem in detecting mixed infection with PCR that was discussed previously must also be considered. There might be more mixed infection than those presented by the results, hence altering the picture. The question that arises is whether this shifting of parasite populations was caused by a new infection during the dry season, or if it can be explained by that
only one parasite population was present in the peripheral circulation at the time of sampling, in a possible mixed infection? These are two probable explanations that have been presented earlier (Hamad et al., 2002; Färnert et al., 1997). One of the hypotheses is that it is possible that just one parasite population is detectable in the peripheral circulation at a given day, while the other parasite populations are harboured in other tissues. This means that only one of the parasite populations gets sampled (Färnert et al., 1997). Furthermore, it remains an open question whether it can be the result of new infections, since it appears to be a complete absence of Anopheles mosquitos during some months of dry season (Hamad et al., 2002). The overall findings in this project are not enough to make any safe conclusions. Hopefully the remaining gaps will bee filled in the future, contributing to these findings to give a larger picture of the characteristics of P.falciparum in eastern Sudan, and to the evolution of drug resistance in the area.
ACKNOWLEDGMENTS
I want to thank Amani Kheir for letting me be part of her project, having confidence in me, and giving me free wings when I has ready. I want also to thank all the people in Göte Swedberg´s group for making me feel very welcome. Especially Hasanthi Karunasekera for sharing her knowledge, giving me inspiration and being a good friend. I would also like to thank my classmates Katarina Järnevi and Rebecka Dahlfors for helping me in the process of writing the report. Last, but not least, I would like to thank Göte Swedberg and Pia Ek.
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