• 1

    Ajao OG, 1978. Malaria and post-operative fever. J Trop Med Hyg 81 :153–155.

  • 2

    Bruce-Chwatt LJ, 1972. Blood transfusion and tropical disease. Trop Dis Bull 69 :825–862.

  • 3

    Manson-Bahr PEC, Ball DR, 1987. Manson’s Tropical Diseases. 19th edition. London: Ballière-Tindall.

  • 4

    Neva FA, Brown HW, 1994. Basic Clinical Parasitology. Sixth edition. New York: Appleton and Lange.

  • 5

    Ali MS, Kadaru AGMY, Mustafa MS, 2004. Screening blood donors for malaria parasite in Sudan. Ethiop J Health Dev 18 :70–74.

  • 6

    Mollison PL, 1994. Blood Transfusion in Clinical Medicine. Ninth edition. Oxford, United Kingdom: Blackwell.

  • 7

    Dover AS, Schultz MG, 1971. Transfusion-induced malaria. Transfusion 11 :353–357.

  • 8

    Talib VH, Khurana SK, 1996. Haematology for Students. Complications of Blood Transfusion. Karachi, Pakistan.

  • 9

    Ali MS, Yousif AG, Mustafa MS, Ibrahim MS, 2005. Evaluation of laboratory procedures applied for malaria parasite screening of Sudanese blood donors. Clin Lab Sci 18 :69–73.

    • Search Google Scholar
    • Export Citation
  • 10

    Schneider J, 1963. Chemoprophylaxis of tropical diseases transmitted by blood transfusion. Transfusion 6 :171.

  • 11

    Cheesbrough M, 1987. Medical Laboratory Manual for Tropical Countries. Second edition. Oxford, United Kingdom: Butter-worth.

  • 12

    Dacie JV, Lewis SM, 1984. Practical Haematology. Sixth edition. Edinburgh, United Kingdom: Churchill Livingstone.

  • 13

    Gramiccia G, 1964. Lessons learned during the final stages of malaria eradication in Europe. Riv Parassitol 25 :157.

  • 14

    Bruce-Chwatt LJ, 1982. Transfusion malaria revisited. Trop Dis Bull 79 :827–840.

  • 15

    Rosenberg R, Wirtz RA, Schneider I, Burge R, 1990. An estimation of the number of malaria sporozoites ejected by feeding mosquito. Trans R Soc Trop Med Hyg 84 :209–212.

    • Search Google Scholar
    • Export Citation

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

IN VITRO PROCESSING OF DONOR BLOOD WITH SULFADOXINE-PYRIMETHAMINE FOR ERADICATION OF TRANSFUSION-INDUCED MALARIA

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  • 1 Department of Haematology, Faculty of Medical Laboratory Sciences, Al Neelain University, Khartoum, Sudan; Medicine, University of Khartoum, Khartoum, Sudan

Transfusion-induced malaria is a problem because of the large number of parasites infused and the weakness of transfused patients. Screening of blood donors or treatment of transfused patients or prospective donors does not eliminate this hazard. It is essential to kill the parasite in vitro in the blood of donors before transfusion. A total of 4,484 blood donors were screened for malaria parasite microscopically using the Giemsa staining technique. Only 30 matched the inclusion criteria of this study. Blood samples were divided into four equal samples. Three concentrations of sulfadoxine-pyremthamine (SP) were added to 90 specimens, and none was added to 30 specimens (controls). Blood specimens were then tested by parasitic, biochemical, and hematologic techniques on the day of collection and after 24 and 48 hours of storage in a blood bank refrigerator. The reduction of malaria parasites was proportional to the concentrations of SP and to the storage period. Blood samples without SP had steady number of the parasites. The lethal dose of SP (the dose that killed 99% of the malaria parasites within 24 hours) was 179.65 μg/L and was highly effective within the 24-hour storage period. This dose did not affect constituents of the stored blood. Thus, for eradication of transfusion-induced malaria by in vitro processing of donors blood, SP is a safe and effective drug. It is recommended that optimal doses of SP be added to donated blood prior to phlebotomy.

INTRODUCTION

Although blood transfusions save many lives, the hazards related to the transmission of disease from the donor to the recipient are significant and well known.1 Many communicable diseases could in principle be transmitted by blood and this results in a formidable list of illnesses that may disqualify many prospective donors.2

Malaria can be transmitted by the inoculation of blood from an infected donor.3 Transfusion-induced Plasmodium falciparum malaria has increased in recent years, probably because the parasite has become increasingly resistant to many drugs.4 Large numbers of parasites can be transmitted through this route of infection. Most patients in need of a blood transfusion are usually weakened by severe disease. Malaria behaves very aggressively in such patients with a higher risk of complications and fatalities.5

Infections with P. falciparum usually decrease within one year, those with P. vivax and P. ovale usually decrease within three years, but those with P. malariae can persist for as long as 50 years. Plasmodium vivax and P. falciparum are the most common species that cause malaria. Plasmodium malariae is also important because of its chronicity and the difficulty in detecting it.3

During the first third of the 20th century, syphilis was a serious medical problem. However, with the introduction of blood banks, syphilis transmission by blood has become a rarity and does not constitute a serious problem because the spirochetes do not survive longer than four days at refrigerator temperature.6 Transmission of malaria through blood transfusions is a greater threat7 because of the capability of human plasmodia to survive in stored blood and even in frozen blood.8

In Sudan, the rate of infection with malaria parasites among blood donors has been estimated by polymerase chain reaction (PCR) to be 21%. This infectivity rate is considerable and many Sudanese patients are at a higher risk of transfusional malaria complications and fatalities.9 This risk is worsened by the fact that the elimination of parasites from blood in vivo, before donation, takes at least 48 hours.10 Therefore, the practical difficulties of this procedure and its unreliability have been reported.2

Systemic screening of possible donors is not a practical solution in malaria-endemic countries because parasitemia in blood donors is often too low to be detected by microscopy or antigen-detection techniques.2 Even when advanced techniques are used, several thousand parasites might be present in one unit of blood and still remain undetected.11 Furthermore, it is neither ethical nor desirable to transfuse infected blood into weak, ill patients because this undoubtedly aggravates their condition. Malaria caused by blood transfusion may also be difficult to control even when optimal doses of antimalarial drugs are used.

These facts necessitate the need for in vitro processing of donor blood that could provide fast and reliable results. Thus, we tested the effect of various concentrations of SP when added to donor blood. These concentrations of SP in a unit of transfused blood are much lower than those given to donors or patients. Research activities for assessment of antimalarial resistance conducted by Department of Biochemistry at the University of Khartoum concluded that chloroquine resistance is increasing, quinine resistance is emerging, but that SP is still fully effective (Khalil IF, unpublished data). The purpose of this study was to determine the lethal dose of SP that eliminates malaria parasites in vitro and the effects of this dose on stored blood.

MATERIALS AND METHODS

The study was conducted at Ahmed Gasim Hospital in Khartoum, Sudan, and was reviewed and approved by Al-Neelain University. The concentration of pyrimethamine was used in the calculations of the three concentrations of SP used. A solution of SP (Fansidar®; F. Hoffmann LaRoche, Basel, Switzerland) (2.5 mL) that contained 25 mg of pyrimethamine was used as follows: 0.25 mL (2.5 mg), 0.5 mL (5 mg), and 0.75 mL (7.5 mg) were diluted in normal saline to obtain 25 mL of three mixtures with concentrations of 100, 200, and 300 mg/L, respectively. Fifty microliters of each concentration were dispensed into separate 50-mL blood unit bags to obtain final pyrimethamine concentrations of 100, 200, and 300 μg/L.

Sample collection took place between October 2002 and January 2004. All individuals who donated blood in Khartoum state hospitals during the study period were included in the studied population if they satisfied the following criteria: presence of malaria parasites in blood films; parasitemia between 1,000 and 80,000 parasites/μL (only asexual stages were considered); growth of parasites on the first day when cultured; and donors did not take quinine in the past 7 days, chloroquine in the past 28 days, and SP in the past 14 days.

One unit (200 mL containing 28 mL of citrate phosphate dextron adenine-1) of malaria parasite–infected donor blood was collected and divided into four equal samples (50 mL) using a blood bank mixer (Biomixer 323; Abelko Innovation, Luleå, Sweden). Three of the samples received the appropriate drug dilution and the other received no drug (control). All blood specimens were tested for parasite culture, platelets count, total leukocyte count, packed cell volume (PCV), percent lysis, osmotic fragility, prothrombin time, activated partial thromboplastin time, serum sodium and potassium levels simultaneously on the day of collection. The blood was then stored in the blood bank refrigerator (4–6°C) and tested after 24 and 48 hours by the same laboratory procedures.

Microscopic identification of malaria parasites was performed as described by Cheesbrough.11 The absolute number of parasites per microliter was determined in thick blood films by counting the number of parasites against 200 white blood cells, multiplying by the total leukocyte count, and dividing by 200. In vitro cultivation of erythrocytic stages of P. falciparum to assess parasite response to SP and confirm the viability or the death of the parasites was conducted using RPMI 1640 medium (Invitrogen, Carlsbad, CA). The minimum inhibitory concentration was determined by computerized probit/log dose response analysis (SPSS, Inc., Chicago, IL). Formation of schizonts at a concentration of 300/≥ 3.75 pmol of SP confirmed resistance.

For counting platelets and white blood cells, blood samples were diluted 1:20 in 1% ammonium oxalate and 2% glacial acetic acid consecutively as described by Cheesbrough.11 Partial thromboplastin time, a screening test for the intrinsic clotting system, i.e., factors XII, XI, IX, VIII, X, and V, prothrombin, and fibrinogen, and prothrombin time, a screening test for the extrinsic clotting system, i.e., factors VII, X, and V, prothrombin, and fibrinogen, were conducted. These tests were conducted as described by Dacie and Lewis12 using a commercial reagent (DiaMed Company, Turnhout, Belgium). The osmotic fragility test, a measure of the surface area/volume ratio of erythrocytes and the effect of various drug doses on erythrocytes, was conducted as described by Dacie and Lewis.12 Packed cell volume, the proportion of whole blood occupied by erythrocytes, was measured by the micro-hematocrit method described by Dacie and Lewis.12

Percentage lysis is an indicator of survival of erythrocytes during the storage period. Five milliliters of each blood sample just after collection were centrifuged for 5 minutes at 12,000 × g. The supernatant was stored at 4°C and used as a control. Five hundred microliters of the centrifuged erythrocytes were diluted in 5 mL of distilled water, stored at 4°C, and used as standard. Both solutions were used in the determination of percentage lysis for the same sample throughout the storage period. One milliliter of each blood sample after 24 and 48 hours was also centrifuged for 5 minutes at 12,000 × g and the supernatant (plasma) of each was tested for hemolysis at a wavelength of 540 nm. Percentage lysis was calculated by dividing the optical density of the tested supernatant by the optical density of the standard and multiplying by 100.

Serum levels of potassium and sodium were measured by the flame photometry using a 410 flame photometer (Sherwood Scientific, Cambridge, United Kingdom).

Data were analyzed using SPSS software (SPSS, Inc.). Non-parametric tests were used for abnormally distributed data.

RESULTS

The number of individuals who donated blood was 4,956; however, 472 (9.5%) were rejected for the following reasons: 303 (6.1%) were infected with hepatitis B virus and 169 (3.4%) were infected with human immunodeficiency virus. The 4,484 collected blood bags were microscopically screened for malaria parasites using the standard Giemsa staining technique. A total of 278 (6.2%) blood bag specimens contained malaria parasites. The parasite densities were < 1,000 in 215 (77%) specimens and ≥ 1,000 in 63 (22.7%) specimens. Of these, 30 (10.8%) samples were resistant to chloroquine and three (1.08%) additional samples showed no growth on the first day when cultured. Thus, the number of blood samples was reduced according to the above-mentioned parameters to 30. All of the accepted blood bags were from asymptomatic male donors between 25 and 35 years of age.

The reduction in malaria parasites when different concentrations of SP were added to donor blood and stored for 48 hours is shown in Table 1. The control sample of donor blood (without SP) showed stable numbers of parasites even after storage for 48 hours. Parasite counts decreased with increasing concentrations of SP and a longer storage period. Parasites were not detectable after 24 hours in blood samples with higher concentrations of SP (200 and 300 μg/L) and after 48 hours in samples containing the minimum concentration (100 μg/L) of SP.

The lethal doses of SP are shown in Table 2. These doses were high when donor blood was stored for 24 hours and lower in blood samples stored for a longer time. Lethal doses also gradually increased when a large number of malaria parasites (99%) were killed (LD99) and decreased when lower numbers of the parasites (50%) were killed (LD50).

Prothrombin time and partial thromboplastin time increased with an increase in SP dose and longer storage period (Table 3). They reached a peak in samples with SP concentrations of 200 and 300 μg/L. The minimum increase was observed with the minimum concentration of SP. Although there was a statistically significant difference between the results among different blood samples stored for the same time period (P < 0.05), the upper levels were within the normal range.

Mean and SD osmotic fragility, which showed increases in samples with higher concentrations of SP and longer storage times, are shown in Table 4. The mean osmotic fragility values reached a peak mainly after storage for 48 hours. No statistically significant difference was observed between the results of the control and the drug-treated blood samples when stored for 24 hours except in samples with a final SP concentration of 300 μg/L. When blood samples were stored for 48 hours, no difference was observed between drug-treated blood samples and the control in samples containing 100 μg/L of SP. Differences were observed in samples containing 200 and 300 μg/L of SP; however, the upper levels were within the normal range.

Mean and SD of PCVs decreased when the time of storage was longer and was independent of the increase in SP dose (Table 5). Samples with a final concentration of 200 or 300 μg/L of SP had significantly different PCVs compared with the control when stored for 24 or 48 hours. However, this statistical difference was within the normal range.

An increased storage time decreased white blood cell and platelets counts (Table 6). The number of platelets correlated with the dose concentration adverse to white blood cell counts. No statistically significant difference was observed between the results of the control and drug-treated blood samples.

Percentage lysis increased in samples containing higher doses of SP and in samples stored for a longer period (Table 7). A statistically significant difference was observed between the control and blood samples containing the highest dose of SP (300 μg/L), but no statistically significant difference was observed between the control and drug-treated blood samples. The amount of lysis observed in these blood samples was insignificant.

The mean and SD serum sodium levels increased with longer storage time and were independent of the drug concentrations (Table 8). A statistically significant difference was observed between the control and blood samples at all three drug doses. Serum potassium levels also increased at high drug concentrations and at longer storage times. After storage for 24 hours, a statistically significant difference (P < 0.05) was observed between the results of the control and blood samples containing 100 and 200 μg/L of SP. Conversely, after 48 hours, no association was observed between results of the control and all blood samples containing 300 μg/L of SP. The upper levels of both electrolytes were both within the normal range for stored blood.

DISCUSSION

Human plasmodia of all species remain infectious in blood stored for at least 1–12 days and most published cases of transfusion-induced malaria occurred within that period. Such cases have also been described with blood stored for two weeks,2 and there is no reason to believe that blood stored up to its shelf life (35 days) is completely safe, although its potential infectivity may be decreased. This issue is probably valid for frozen blood cell and platelets, but not freeze-dried concentrates.8

Direct microscopic examination of donor blood is of little value. Such a time-consuming procedure is obviously impractical, especially with regard to asymptomatic blood donors who often have low parasite densities at the submicroscopic level.13 Detection of malaria antibodies provides evidence of an immune response to current or past infection. However, these test results may remain positive for more than 10 years after the parasitemia has resolved. Therefore, detection of malaria antibodies to screen blood donations would result in the exclusion of otherwise healthy persons.14

Although the PCR has an increased sensitivity compared with blood film examination, several thousand malaria parasites might be present in a unit of blood (450 mL) and still not be detected.11 This number of parasites is much larger compared with the 12–20 sporozoites delivered by the mosquito bite.15 This indicates the ineffectiveness of testing blood donors to eliminate the risk of malaria transmission by blood transfusion, which is consistent with the failure of the screening system used by blood banks in the United States to prevent the occurrence of such cases.7

In the present study, the number of parasites and parasites densities in blood samples treated with three doses of SP showed a significant difference when stored for 24 hours. Nevertheless, they were insignificantly different when blood samples were stored for 48 hours. This is likely due to effectiveness of the different concentrations of SP, which was reflected by the effect of even low concentrations of this drug.

The lethal dose of SP (179.65 μg/L) was greater than that reported by Bruce-Chwatt in 1985 (10–100 μg/L). This increase is probably due to the behavior of the parasite towards the drug over the previous 19 years, which has resulted in adaptation of the parasite to become relatively resistant. Also, the difference in the strains may explain this increase.

The effect of SP on all constituents of stored blood was assessed to ensure whether the doses of drugs were safe. The mean prothrombin time and partial thromboplastin time, which are indicators of the clotting system, increased in samples containing higher drug concentrations. However, this increase also showed a correlation with the time of storage. Even at the higher drug dose, the increase in these times was acceptable. Thus, the lethal dose of the drug does significantly affect plasma coagulation factors.

The osmotic fragility test, which measures the functions and integrity of the erythrocyte membrane, was used to measure the effect of SP on the malaria parasite and the erythrocyte membrane. Mean osmotic fragility values were significantly correlated with the storage time; they increased when blood samples were stored for 48 hours compared with those stored for 24 hours. However, lethal doses of the drug had no effects on the flexibility of the erythrocyte membrane.

The hematocrit is an indication of shrinking or swelling of red blood cells. In stored blood, hematocrit values are decreased because of dilution by the anticoagulant solution. Some investigators have described a relative increase in hematocrit values of stored blood with an increase in storage time due to the entry of plasma fluids into the erythrocytes during storage.6 In the present study, PCV values decreased proportionally with the length of the storage period. However, SP did not significantly affect these values. The red blood cells that hemolyzed during storage may explain the decrease in PCV in the present study.

We also studied the effect of SP on platelet counts. This study showed a significant decrease in platelet counts that correlated with increasing concentration of SP. However, the effect of lethal doses of SP on platelet counts appears to be acceptable; counts were within the reference values for stored blood.

Percent lysis in stored blood reflects the unfavorable effect of storage or applied drugs on the viability of erythrocytes. This value in the present study increased in proportional to the concentrations of the drug as well as the storage period. Although the highest value of lysis was 0.22%, it was low compared with that reported in normal stored blood (1%) by Mollison.6

Storage of donor blood results in an increase in both sodium and potassium levels, which is likely to be due to inactivation of active transport of these electrolytes across the red blood cell membrane. Blood samples showed a poor correlation between sodium levels and drug concentration, despite a good correlation with the storage period. However, these findings were not observed for potassium levels. They did not show a correlation with the drug concentration, but did show a correlation with the duration of storage. However, the increase in the potassium level was acceptable when blood samples were treated with the lethal dose of SP.

We have shown that in vitro treatment of donor blood with SP is simple and applicable because the concentrations of SP used can be safely added to the constituents of stored blood (anticoagulant and preservatives) without incompatible interactions. Moreover, this process is inexpensive because one ampule of SP can be used for 250 blood units (bags). We recommend that SP be added to stored blood to give a final concentration of 180 μg/L. Future studies are recommended to select highly effective and safe antimalarial drugs based on patterns of resistance and side effects and to assess the safety of this process.

Table 1

Positivity and mean values of malaria parasite counts during storage at 2–8°C*

Day of collectionTwenty-four hoursForty-eight hours
SamplePositivityMeanPositivityMeanPositivityMean
* Mean values are in parasites/μL.
F1 = 100 μg/L of sulfadoxine-pyrimethamine (SP).
F2 = 200 μg/L of SP.
F3 = 300 μg/L of SP.
F130 (100%)3,2735 (16.7%)69.32 (6.7%)11
F230 (100%)3,2732 (6.7%)300 (0%)0
F330 (100%)3,2730 (0%)00 (0%)0
Control30 (100%)3,27330 (100%)3,27330 (100%)3,273
Table 2

Lethal doses (LDs) of fansidar that kill 50%, 95%, and 99% of the parasites in blood samples stored for 24 and 48 hours in the blood bank (2–8°C)

Lethal doses
Storage periodLD50LD95LD99
Twenty-four hours60.68 μg/L144.80 μg/L179.65 μg/L
Forty-eight hours52.95 μg/L80.16 μg/L91.43 μg/L
Table 3

Prothrombin time (PT) and activated partial thromboplastin time (APTT) in blood samples stored at 2–8°C*

First day24-hr incubation48-hr incubation
SamplePTAPTTPTAPTTPTAPTT
* Times are in seconds. For definitions of samples, see Table 1.
F115.6736.2016.1730.0318.2041.33
F215.6736.2016.9340.9019.2042.03
F315.6736.2017.1042.5019.9044.90
Control15.6736.2015.9338.40017.9740.867
Table 4

Osmotic fragility values of blood samples stored at 2–8°C*

First day24-hr storage48-hr storage
SampleMeanSDMeanSDMeanSD
* Values are percentages (g/dL of NaCl). For definitions of samples, see Table 1.
F10.45100.003050.47500.03410.4800.03373
F20.45100.003050.48500.03970.5150.03256
F30.45100.003050.49000.04980.5050.03560
Control0.45100.003050.46000.0310.4600.03050
Table 5

Haematocrit values of blood samples stored at 2–8°C*

First day24-hr storage48-hr storage
SampleMeanSDMeanSDMeanSD
* For definitions of samples, see Table 1.
F142.702.73139.403.75638.506.224
F242.702.73135.502.85036.105.155
F342.702.73135.905.05435.504.711
Control42.702.73140.503.97239.507.960
Table 6

White blood cell (WBC) and platelet (PLT) counts in blood samples stored at 2–8°C*

First day24-hr storage48-hr storage
SampleWBCPLTWBCPLTWBCPLT
* Values are cells per microliter. For definitions of samples, see Table 1.
F13.250225.0003.130170.5002.740171.000
F23.250225.0002.820166.1072.460161.000
F33.250225.0003.090161.5002.600158.000
Control3.250225.0002.940208.0003.230178.000
Table 7

Percentage lysis of red blood cells stored at 2–8°C*

First day24-hr storage48-hr storage
SampleMeanSDMeanSDMeanSD
* For definitions of samples, see Table 1.
F1000.140.070.160.08
F2000.160.060.170.06
F3000.170.090.220.07
Control000.090.070.160.1
Table 8

Serum levels of sodium (Na) potassium (K) in blood samples stored at 2–8°C*

First day24-hr storage48-hr storage
SampleNaKNaKNaK
* Values are in mmol/L.
For definitions of samples, see Table 1.
F1140.702.833165.304.3173.24.800
F2140.702.833170.003.967172.64.533
F3140.702.833169.003.8001875.0
Control140.702.833157.903.7154.304.4

*

Address correspondence to Mohamed S. M. Ali, Department of Haematology, Faculty of Medical Laboratory Sciences, Al Neelain University, P.O. Box 12702, Khartoum, Sudan. E-mail: mohdaru@hotmail.com

Authors’ addresses: Mohamed S. M. Ali, Department of Haematology, Faculty of Medical Laboratory Sciences, Al Neelain University, Khartoum, Sudan, E-mail: mohdaru@hotmail.com. Abdul G. M. Y. Kadaru, Faculty of Medicine, University of Khartoum, Khartoum, Sudan.

REFERENCES

  • 1

    Ajao OG, 1978. Malaria and post-operative fever. J Trop Med Hyg 81 :153–155.

  • 2

    Bruce-Chwatt LJ, 1972. Blood transfusion and tropical disease. Trop Dis Bull 69 :825–862.

  • 3

    Manson-Bahr PEC, Ball DR, 1987. Manson’s Tropical Diseases. 19th edition. London: Ballière-Tindall.

  • 4

    Neva FA, Brown HW, 1994. Basic Clinical Parasitology. Sixth edition. New York: Appleton and Lange.

  • 5

    Ali MS, Kadaru AGMY, Mustafa MS, 2004. Screening blood donors for malaria parasite in Sudan. Ethiop J Health Dev 18 :70–74.

  • 6

    Mollison PL, 1994. Blood Transfusion in Clinical Medicine. Ninth edition. Oxford, United Kingdom: Blackwell.

  • 7

    Dover AS, Schultz MG, 1971. Transfusion-induced malaria. Transfusion 11 :353–357.

  • 8

    Talib VH, Khurana SK, 1996. Haematology for Students. Complications of Blood Transfusion. Karachi, Pakistan.

  • 9

    Ali MS, Yousif AG, Mustafa MS, Ibrahim MS, 2005. Evaluation of laboratory procedures applied for malaria parasite screening of Sudanese blood donors. Clin Lab Sci 18 :69–73.

    • Search Google Scholar
    • Export Citation
  • 10

    Schneider J, 1963. Chemoprophylaxis of tropical diseases transmitted by blood transfusion. Transfusion 6 :171.

  • 11

    Cheesbrough M, 1987. Medical Laboratory Manual for Tropical Countries. Second edition. Oxford, United Kingdom: Butter-worth.

  • 12

    Dacie JV, Lewis SM, 1984. Practical Haematology. Sixth edition. Edinburgh, United Kingdom: Churchill Livingstone.

  • 13

    Gramiccia G, 1964. Lessons learned during the final stages of malaria eradication in Europe. Riv Parassitol 25 :157.

  • 14

    Bruce-Chwatt LJ, 1982. Transfusion malaria revisited. Trop Dis Bull 79 :827–840.

  • 15

    Rosenberg R, Wirtz RA, Schneider I, Burge R, 1990. An estimation of the number of malaria sporozoites ejected by feeding mosquito. Trans R Soc Trop Med Hyg 84 :209–212.

    • Search Google Scholar
    • Export Citation
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