Background
In the past decade, remarkable reductions in malaria burden have been achieved largely through widespread access to and use of artemisinin-based combination therapy and insecticide-treated bed nets [
1]. However, since 2015, progress has stalled [
2] and elimination is unlikely to be achieved with these conventional methods alone in most settings in Africa [
3]. One of the biggest challenges to malaria control and elimination is effectively interrupting the efficient process of malaria transmission. New tools or implementation strategies will likely be required that specifically aim to reduce malaria transmission [
4]. In order to effectively target transmission reduction and develop and implement transmission-blocking interventions, a thorough understanding of the dynamics and epidemiology of malaria transmission in different settings is needed.
The transmission of malaria depends on the presence of mature sexual stage parasites (gametocytes) in the peripheral blood. During a blood meal,
Anopheles mosquitoes must imbibe at least one male and one female gametocyte to become infected, but other host or parasite factors may also contribute to this, such a transmission reducing immunity. To understand how likely it is that individuals contribute to onward transmission, assays are needed for accurate assessment of gametocyte infectivity to mosquitoes. This is commonly measured by direct membrane feeding assays (DMFAs) where venous blood is fed to mosquitoes, or direct skin feeding assays, where mosquitoes feed directly on the skin of the volunteer [
5‐
11]. In DMFAs, venous blood collected from malaria infected individuals is offered to female
Anopheles mosquitoes using water-jacketed glass feeders. These feeders are connected to a circulating water bath to maintain gametocyte temperature during feeding. Maintaining gametocyte temperature is important since a drop in temperature may result in gametocyte activation [
12] while high temperatures (~ 40 °C or higher) may inactivate gametocytes and gametes [
13‐
16].
Blood drawn for DMFAs must therefore be fed to mosquitoes as quickly as possible after collection to eliminate any detrimental effect of temperature change [
17]. This can be logistically challenging in the field where participants in DMFA studies may be recruited far from the insectary facilities, resulting in either lengthy travel for volunteers, or restricting the available population for infectivity assessment to those near the facilities. To overcome these limitations, a method to store and transport gametocyte infected blood from the field to the laboratory and prevent premature activation or inactivation of gametocytes before mosquito feeding is needed.
In this study, the temperature range for storage of gametocyte infected blood to maintain gametocyte infectivity is determined, and a simple, cheap, field applicable method for collecting and transporting blood is presented.
Methods
Mosquito rearing
Anopheles stephensi Sind-Kasur Nijmegen strain [
18] (Radboudumc) were reared at 30 °C, and
Anopheles coluzzii (Burkina Faso and The Gambia) were reared at 27 °C. All were kept at ~ 70–80% relative humidity (RH) on a 12-h day/night cycle and were fed on 5–10% glucose. Female mosquitoes between 2 and 6 days post emergence were used for all mosquito feeding assays. After feeding on gametocyte infected blood, mosquitoes were stored either at 26 °C (Radboudumc) or 27 °C (Burkina Faso and The Gambia).
Standard membrane feeding assays (SMFAs)
SMFAs were performed at Radboudumc, Nijmegen, The Netherlands, using cultured
Plasmodium falciparum gametocytes as previously described [
19]. In brief,
P. falciparum was cultured at 37 °C in an automated incubator under a continuous gas flow of 4% CO
2, 3% O
2 and 93% N
2. Gametocytes were produced from asynchronous cultures (day 0, 1% parasitaemia and 5% red blood cells), which were harvested at day 15. Gametocyte infected blood meals were prepared by adding 900 µl mature stage 5 gametocyte culture to 600 µl packed red blood cells, this was briefly centrifuged, the supernatant removed and 600 µl human serum added. For the long-term storage experiments, gametocyte infected blood meals were either fed immediately to mosquitoes or were stored for 4 h in a tube in a thermos flask filled with water at a range temperatures, measured using a calibrated probe thermometer accurate to ± 0.2 °C (Traceable® Ultra digital thermometer, VWR 620-2079) before feeding to mosquitoes. The water temperatures chosen were either the temperature hypothesized to be the optimal gametocyte storage temperature (the temperature of the human body), 37 °C (range: 36.9–37.2 °C) or an arbitrarily selected lower (35.5 °C [range: 35.4–35.6 °C]) or higher (37.8 °C [range: 37.8–37.8 °C]) temperature. Thermos flasks (Stainless king food flask 470 ml, Thermos) were stored either at room temperature (RT; mean 21.3 °C, range: 20.4–22.1 °C) or in an incubator set at 32 °C (range: 31.9–32.2 °C) or 42 °C (range: 41.4–42.1 °C). These storage temperatures were selected to reflect the range of temperatures that might be experienced during thermos flask storage in a malaria endemic county. The exact thermos storage temperature and water temperatures are presented in Additional file
1: Table S1. For the short-term storage experiments, gametocyte infected blood meals were stored for 15 min at RT or in a heat block at a range of temperatures (30 °C, 35.5 °C, 38 °C, 40 °C or 42 °C) before feeding to mosquitoes. For all SMFAs, female mosquitoes were offered the infected blood meal in glass mini-feeders (300 μL) attached to a circulating water bath set at 39 °C and allowed to feed in the dark for 15 min [
19]. A total of 30–60 mosquitoes in 2 or 3 cups were used for each experimental condition. Mosquitoes were maintained on 5% glucose and dissected 6–8 days after feeding and the number of developed oocysts per mosquito was determined by microscopy after 1% mercurochrome midgut staining.
Ethical approval
The Burkina Faso study was approved by the London School of Hygiene and Tropical Medicine ethics committee (Review number: 14724), the Centre National de Recherche et de Formation sur le Paludisme institutional review board (Deliberation N° 2018/000,002/MS/SG/CNRFP/CIB) and the Ethics Committee for Health Research in Burkina Faso (Deliberation N° 2018–01-010). The Gambia study was approved by the London School of Hygiene and Tropical Medicine ethics committee (Review number:15993) and by The Gambia Government/MRC Joint Ethics Committee (Review number:1621). All participants gave informed consent before inclusion in the studies.
Gametocyte positive blood samples
Gametocyte positive human blood samples were obtained from individuals as part of ongoing studies in Burkina Faso and The Gambia. In Burkina Faso, participants were microscopy-positive gametocyte carriers aged 10–15 years old, recruited from screening campaigns in the Saponé health and demographic surveillance system area, ~ 45 kms Southwest of Ouagadougou. In The Gambia, participants were microscopy-positive gametocyte carriers aged > 2 years, passively recruited from four health facilities (Basse Hospital, Sabi, Sotuma Sere, Gambissara) in the South Bank of the Upper River Region in The Gambia.
Direct membrane feeding assays (DMFAs)
Thermos flask water temperature setup
Thermos flasks (Stainless king food flask 470 ml, Thermos) were filled (to 2 cm below the top of the flask) with water between 35.5 and 36 °C, measured with a calibrated probe thermometer (accurate to ± 0.2 °C, Traceable® Ultra digital thermometer, VWR 620-2079) and immediately sealed. Gametocyte infected blood samples were collected in Lithium heparin vacutainers and either fed to mosquitoes immediately (control) or transferred to the thermos flask immediately after phlebotomy. One thermos flask was used per blood donor to avoid repeated opening of the flask, and 2 tubes of blood were collected from each donor. Thermos flasks were used within 1 h of it being filled with water. Thermos flasks were then stored at ambient temperature in the laboratory or in an air-conditioned car (range: 25.5–29 °C) for a maximum of 4 h from the time of filling and the stored blood sample was then used for mosquito feeding.
Feeding procedures
For all DMFAs, female mosquitoes were offered the infected blood meal in glass mini-feeders (300 μL) attached to a circulating water bath at 37–39 °C and allowed to feed in the dark for 15–20 min. A total of 3 cups of 30 mosquitoes were used for all experimental conditions in The Gambia, and 2 cups of 60 mosquitoes were used for all experimental conditions in Burkina Faso. Mosquitoes were maintained on 5–10% glucose until 7–8 days after feeding when they were dissected and the number of developed oocysts per mosquito was determined by microscopy after 0.5% mercurochrome midgut staining [
20].
Statistical analysis
Statistical analysis was performed using GraphPad Prism (ver. 8). Mosquito infectivity in SMFAs was analysed by comparing groups with the Kruskal–Wallis test comparing each experimental condition to the control with Dunns multiple comparison test. Mosquito infectivity in DMFAs was analysed by comparing oocyst intensity (average oocysts per mosquito) or prevalence of infection (% of infected mosquitoes) by Wilcoxon matched-pairs signed rank test. Agreement between the immediate DMFAs or DMFAs with stored gametocytes was analysed using Spearmans correlation of % mosquitoes infected before and after gametocyte storage and Bland–Altman analysis to visualize and test whether systematic bias in infection rates occurred between the two methods.
Discussion
In this study, a method to maintain P. falciparum gametocyte infectivity during blood collection in the field and transportation to the insectary for mosquito feeding assays is presented. The optimal storage conditions preventing gametocyte activation or inactivation were first assessed with cultured gametocytes and then confirmed using natural gametocyte carriers from two malaria endemic settings. The findings corroborate that temperature can play a key role in maintaining the infectivity of P. falciparum gametocytes. Infectivity was not affected when gametocytes were stored in thermos flasks in water at 35.5 °C for up to 4 h and the thermos flask was stored at ambient temperatures ranging from 21.3 to 32 °C.
DMFAs are more commonly used than direct skin feeding assays to assess the infectiousness of gametocyte carriers to mosquitoes, largely because they minimize the discomfort experienced by volunteers, which is particularly important when sampling young children. DMFAs may, therefore, be more readily acceptable by both local communities and ethics committees [
21], especially when repeated assessments of infectivity are made. The relationship between gametocyte density and infection success in mosquitoes has been well studied, but differs slightly by settings [
11,
21,
22]. These differences could plausibly be due to the different populations, different levels of malaria exposure and resulting levels of transmission-blocking immunity, different mosquito species, or parasite genotypes [
8,
9,
11]. However, for reliable assessments of infectivity by DMFA it is crucial to minimize technical differences in how the assays are performed between settings [
21,
22]. Here, evidence is presented that variation in blood storage temperature and duration of storage, as well as feeder temperature, could have a significant impact on transmission.
The results show that the optimal temperature for longer-term storage (4 h) is 35.5 °C. The maintenance of gametocyte infectivity at this temperature is consistent with previous studies which showed a drop in temperature of at least 5 °C from the standard 37 °C in the human body, is required to activate
P. falciparum gametocytes [
12]. Also consistent with previous data [
13,
14,
16,
23], our results show that as little as 15 min (mimicking the time the gametocytes are present in the feeder) at 42 °C is sufficient to inactivate gametocytes, with transmission being almost completely prevented. Taken together, these data suggest that the temperature in the feeders should not exceed 40 °C during DMFAs.
With the 4-h storage experiments, not only temperatures above 40 °C but also lower temperatures appeared to reduce gametocyte infectivity. It was surprising to find that storing the gametocytes in the thermos flask in water at 37 °C for 4 h was associated with reduced transmission efficiency in some experiments. This was seen most often when the ambient temperature was high (i.e. 32 °C or 42 °C), and not when it was the typical room temperature in an air-conditioned European laboratory (~ 21.3 °C). The temperature of 37 °C would be hypothesized to be ideal for gametocytes, as it mimics their natural environment in the body, and previous studies have indeed shown that
P. falciparum gametocyte activation in vitro was prevented when they were maintained at 37 °C for 1 h [
24]. In agreement with this, short-term storage for 15 min at temperatures up to 40 °C in the study presented here did not reduce transmission. This suggests that either the duration of exposure to high temperatures, or the fact that the gametocytes are stored in venous blood collected in lithium heparin anticoagulant, or both, may also be important factors for gametocyte inactivation, although this is not evaluated here and warrants further investigation. Similarly, the increase in transmission efficiency following 4-h storage compared to immediate feeding, as observed in some experiments, also requires further study. Factors such as the presence of anticoagulants during storage and the duration of storage permitting continued gametocyte maturation may play a role.
Altogether, the data demonstrates that temperature fluctuations influence gametocyte infectivity, seen most acutely with higher temperatures, including high ambient temperatures. Temperature should thus be carefully controlled when collecting and transporting gametocyte infected blood in regions where ambient temperatures could reach over 40 °C. An effect of temperature on gametocyte infectivity has been seen before, however, the current study adds value with the large number of replicates in the SMFA evaluating varying thermos and ambient temperatures. This allowed an informed decision on the optimal temperature conditions for use in DMFA experiments using natural gametocytes carriers in the field.
Conclusion
This study presents the validation of a method to maintain P. falciparum gametocyte infectivity during blood collection and transportation for DMFAs. The optimal conditions (storage in a thermos flask in water at 35.5 °C for up to 4 h with the thermos flask stored between 21.3 and 32 °C) were determined in SMFAs and verified using gametocyte infected venous blood samples from natural gametocyte carriers. With the proposed approach, samples can be transported from more remote settings to the insectary within 4-h without affecting gametocyte infectivity and thus maintaining assay quality. This method will facilitate widespread, accurate assessment of malaria transmission dynamics in the field and this knowledge will contribute to malaria control strategies as progress is made towards elimination.
Acknowledgements
The authors thank Jolanda Klaassen, Laura Pelser-Posthumus, Astrid Pouwelsen and Jacqueline Kuhnen for mosquito rearing and mosquito membrane feeding experiments at Radboudumc; Traore Ben Idriss, Damiba Wilfried, Nabolle Madi and Traore Mamoudou for mosquito rearing and dissection in Burkina Faso; the mosquito rearing and dissection team in The Gambia; the study nurses, lab technicians, drivers and clinic leadership in all the health districts. Special thanks to the study volunteers for their participation and the communities for their support.
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