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ABSTRACT |
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INTRODUCTION |
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Seventy-five percent of Ethiopia is malarious, and 65% of its population lives in this area and is at risk for malaria. Five to six million clinical cases and more than 600,000 confirmed cases have been reported from health facilities in non-epidemic years. This indicates only a portion of the actual magnitude of this problem since there is a poor accessibility to and use of health services. Plasmodium falciparum is the predominant species that causes severe and complicated clinical manifestations and almost all malaria deaths.2
Effective treatment is essential for malaria control. However, drug-resistant malaria has become a challenge in malaria control programs in recent years. The emergence of multidrug-resistant strains of P. falciparum has compromised the effectiveness of routinely used anti-malaria drugs. This has threatened the use of the cheap and safe drugs in resource-poor countries where the mosquito control has been ineffective.
Sulfadoxine/pyrimethamine (SP) has been used as an affordable alternative treatment of uncomplicated malaria cases in chloroquine-resistant areas of Africa.3 However, in some east African countries, including Ethiopia, SP was adopted earlier and has been used extensively as a first-line treatment due to the high prevalence of chloroquine-resistant strains of P. falciparum. This antifolate drug was used intensively in these areas, which has led to the selection of resistant strains against the drug.4
Resistance to pyrimethamine is associated with mutations in the gene encoding the parasite enzyme dihydrofolate reductase (DHFR), and resistance to sulfadoxine is correlated with mutations in the parasite gene for dihydropteroate synthetase (DHAP).5,6 The level of resistance is associated with the number of mutations in the genes for these two enzymes. Therefore, multiple mutations in the two genes are considered to be responsible for SP treatment failure.7
Mutations in DHFR have been reported at codons 16, 51, 59, 108, and 164 in a number of geographic isolates.8 Mutations S108 to N108 or T108 in DHFR has been proposed as the main mechanism of resistance against pyrimethamine. All multiple mutations emerge from stepwise selection of a single mutant at position 108 of the DHFR gene. It has also been shown that the resistance level is significantly increased by additional sequence changes at positions 51 (N51 to I51) and 59 (C59 to R59) in DHFR.8,9 In addition, a point mutation at codon 164 has been suggested to be responsible for the development of resistance to chlorproguanil-dapsone.10
The major amino acid mutation in the DHPS gene is at residue 437 (A to G), which plays a major role in the development of clinical resistance against sulfadoxine. Mutations in the DHPS gene associated with resistance to sulfadoxine include a change of S436 to F436, A437 to G437, K540 to E540, A581 to G581, and A613 to S613 or T613. Normally, multiple DHPS mutations result in a synergistic effect on SP resistance.9
A molecular geographic survey for the occurrence of P. falciparum drug resistance is of paramount importance for quantifying antimalarial drug efficacy and for monitoring the emergence of drug-resistant malaria.11 Molecular methods have an important application in surveillance programs as attractive tools to detect drug-resistant mutants in epidemiologic surveys. Moreover, molecular tests have many advantages compared with in vitro testing, which requires a complex parasite cultivation technique and several days to perform.12–14
Patients infected with parasites carrying the DHPS G437E540 double mutant and the DHFR S108I51R59 triple mutant had a specifically high relative risk of treatment failure compared with those infected with parasites carrying only the DHFR triple mutant.15 In a similar study, the quintuple mutant (3 DHFR and 2 DHPS mutations) was shown to be a relevant molecular marker of SP treatment failure among uncomplicated P. falciparum malaria patients.7
Since the molecular basis of resistance against the anti-folate drugs is well characterized, it is advisable to use molecular methods to detect drug-resistant malaria. Based on this principle, we have performed a molecular survey on P. falciparum isolates collected from Ethiopia to measure the level of antifolate resistance in this country.
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METHODS |
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Patients with clinically suspected malaria attending the Jimma Health Center (Jimma, Ethiopia) were screened and included in the study. Only those patients with uncomplicated malaria who had P. falciparum infections and who gave informed consent were included in the study. Severely ill patients and those who had a febrile illness other than malaria were excluded from the study. The study was reviewed and approved by the Addis Ababa University Ethical Committee. Individual written informed consent was obtained from each patient.
Thin and thick blood films were prepared and stained with Giemsa for the detection of P. falciparum infection. For mutation analysis, 20 µL of blood was collected by finger prick, blotted in triplicate on a filter paper, and air-dried. The filter papers from patients with malaria were wrapped separately in a plastic bag and transported at room temperature. DNA was isolated using the QIA amp® DNA Mini Kit (Qiagen, Hilden, Germany) following the manufacturer’s instruction.
The samples were analyzed using a nested polymerase chain reaction (PCR) and DNA sequencing to detect variation in the DHFR and DHPS genes. The extracted DNA (2 µL) was used as a template in 50-µL PCR that contained 0.2 µM of each oligonucleotide primer, 1x PCR buffer (Qiagen), 2.5 mM MgCl2, 0.2 mM dNTPs, and 0.02 units/µL of Taq DNA polymerase. Accordingly, a fragment of the DHFR gene containing codons 16, 51, 59, 108, and 164 and a fragment of the DHPS gene containing codons 436, 437, 540, 581, and 613 was amplified by a nested PCR approach.
In the first reaction, a 665-basepair (bp) portion of the DHFR gene was amplified by using primers Amp1 (5'-TTT ATA TTT TCT CCT TTT TAT-3') and Amp22 (5'-TTA CTA GTA TAT ACA TCG CTA ACA G-3'). Similarly, a 727-bp portion of the DHPS gene was amplified by using primers sulf5' (5'-GGT ATT TTT GTT GAA CCT AAA CG-3' and sulf3' (5'-TCC AAT TGT GTG ATT TGT CCA C-3').
For the second reaction of the DHFR gene, 3 µL of amplified material from the first PCR product was added to the second PCR mixture. Primers SP1 (5'-ATG ATG GAA CAA GTC TGC GAC-3') and Amp22 were used to amplify a 646-bp fragment containing codons 16, 51, 59, 108, and 164.
The second round PCR of the DHPS gene was done using similar reaction conditions like those used for DHFR, but two different primer pairs were used to obtain two separate fragments. Primers sulf5' and Leo2' (5'-CTG GAT TAT TTG TAC AAG CAC-3') were used to amplify a 319-bp fragment of the DHPS gene containing the sequence of codons 436 and 437. Similarly a 472-bp DHPS fragment that included codons 540, 581, and 613 was amplified using the primer pair DS-5F (5'-GAA TGT GTT GAT AAT GAT TTA G- 3') and sulf3'.
The products from the nested PCR were subjected to electrophoresis on a 1% agarose gel. Gels were stained with CYBR® GREEN I nucleic acid gel stain (Cambrex Bioscience, East Rutherford, NJ) and visualized on a dark reader transilluminator (Clare Chemical Research, Dolores, CO).
The amplified DNA was purified by a PCR purification kit (EZNA.® Cycle Pure Kit; Peqlab, Erlangen, Germany), following the supplier’s instructions. The DNA was then sequenced by using Big Dye 1.1 (Applied Biosystems, Foster City, CA) and purified by DNA grade SephadexTM (Amersham Biosciences AB, Uppsala, Sweden). A genetic analyzer 3100 (Applied Biosystems) was used for analysis of DNA. The DNA sequences were transferred to the Bio-edit sequence alignment program (http://www.mbio.ncsu.edu/BioEdit/bioedit.html) for detection of point mutations and sequence comparison.
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RESULTS |
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The prevalence rate of previously known mutations has been detected. The sequence profile in this study showed that all isolates had double mutations at residues 51 and 108, as shown in Table 1
(I51N108). Sixty-seven (54%) of the isolates had a triple mutation in the DHFR gene (I51R59N108).
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Higher mutation rates in the DHFR and DHPS genes indicate a higher level of resistance against SP treatment in Ethiopia, similar to previous in vivo studies. Chlorproguanil-dapsone, another antifolate drug, is dependent on the presence of a point mutation at position 164 of the DHFR gene. In this study, as previously reported in other African countries, there is no point mutation at this position in any isolates.
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DISCUSSION |
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In Ethiopia, the development of resistance to currently used anti-malaria drugs has decreased the effectiveness of early diagnosis and treatment of malaria. Sulfadoxine-pyrimethamine was introduced as a first-line drug for treatment of uncomplicated P. falciparum malaria in July 1998. When it was introduced, the treatment failure rate was approximately 5%. In a recent nation-wide study that was conducted from October to December 2003, the mean failure rate had increased to 36% at the 14-day follow-up and 72% on the 28-day follow-up.2
The occurrence of DHFR and DHPS mutations in our study is similar to findings from Malawi, the first African country to adopt SP as first-line treatment of uncomplicated malaria.3 We found that 54% of the samples had quintuple mutations; a higher (78%) prevalence was reported in Malawi. The prevalence of DHPS double mutations (100%) was higher in Ethiopia than that reported in Malawi.3
Quintuple mutations served as an indicator for the presence of SP-resistant strains in Mali, where no quintuple mutation was found.16 A similar analysis in Kenya, which is in the same geographic region as Ethiopia, reported that 10% of the samples had quintuple mutations.15 The result of our study showed a higher rate of resistance against SP. This should warrant that immediate measures have to be taken to identify the most appropriate treatment option. Based on the World Health Organization guideline for Africa, artemether-lumefantrine was adopted as a first-line treatment against uncomplicated P. falciparum malaria in Ethiopia; however, it is too costly and unavailable.
Based on the absence of L164 mutation in the DHFR gene of P. falciparum, chlorproguanil-dapsone was recommended for Africa as affordable, readily available, and effective treatment option against SP-resistant strains of P. falciparum. It is rapidly eliminated from the body, resulting in low selection pressure for drug resistance.16,17 Chlorproguanil-dapsone may be an affordable alternative, but it may cause problems in areas with a high deficiency of glucose-6-phosphate dehydrogenase. It is not known whether in Ethiopia this deficiency exists.18,19
Artemisinin-based combinations have a short shelf-life (two years), and emergency supplies have to be quantified and maintained in well-managed stocks.20 Moreover, WHO has indicated that the rapid increase in demand for ACTs may result in shortages in the production and delivery of the drug because artimisinin compounds are derived from the plant Artemisia annua. The cultivation of this plant, as well as the extraction and the manufacturing process, requires approximately three years. Therefore, agricultural production should be coordinated with the increased demand for pharmaceutical products. WHO forecasts that a massive scale-up of production is needed to meet the global requirements of the drug in 2005.
Additional combination therapies and monotherapies have been recommended as effective treatment alternatives against P. falciparum. However, the ideal (inexpensive, readily available, effective, and safe) regimen for Africa is still not known.
Received May 20, 2005. Accepted for publication July 14, 2005.
Acknowledgments: We thank the Graduate School of Addis Ababa University for organizing the study, Almaz Demissei (Jimma Health Centre) for recruiting patients, and Andrea Weierich and Velia Grummes for technical help at the Sequencing Facility of the Institute for Tropical Medicine.
Financial support: This work was supported by grants to Tamirat Gebru-Woldearegai from the Deutscher Akademischer Austausch Dienst and by the Professor Josef und Erika Hesselbach Stiftung to Jürgen F. J. Kun.
* Address correspondence to Jürgen F. J. Kun, Institute for Tropical Medicine, Department of Parasitology, University of Tübingen, Wilhelmstr. 27, 72074 Tübingen, Germany. E-mail: juergen.kun{at}uni-tuebingen.de 
Authors’ addresses: Tamirat Gebru-Woldearegai, Institute for Tropical Medicine, Department of Parasitology, University of Tübingen, Wilhelmstr. 27, 72074 Tübingen, Germany, Telephone: 49-7071-298-2195, Fax: 49-7071-294-684, E-mail: tamiratgw2002{at}yahoo.com (present address: P.O. Box 70226, Addis Ababa, Ethiopia, Telephone: 251-9-645-665). Asrat Hailu, P.O. Box 28017/1000, Addis Ababa, Ethiopia, Telephone: 251-9-480-993, Fax: 251-1-513099, E-mail: hailu_a2004{at}yahoo.com. Martin P. Grobusch, Institute for Tropical Medicine, Department of Parasitology, University of Tübingen, Wilhelmstr. 27, 72074 Tübingen, Germany, Telephone: 49-7071-298-0234, Fax: 49-7071-294-684 (present address: Faculty of Health Sciences, University of the Witwatersrand, 7 York Road, Parktown 2196, Johannesburg, South Africa, Telephone: 27-11-489-8537, Fax: 27-11-489-8511, E-mail: grobuschm{at}pathology.wits.ac.za). Jürgen F. J. Kun, Institute for Tropical Medicine, Department of Parasitology, University of Tübingen, Wilhelmstr. 27, 72074 Tübingen, Germany, Telephone: 49-7071-298-2191, Fax: 49-7071-294-684, E-mail: juergen.kun{at}uni-tuebingen.de.
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