Fluorescence and Failure

By Jack Sewell

Project aims

This post is about the malaria lab project that we've been working on full-time this semester. To avoid giving away too much information about what the lab is investigating, I won't mention the specific proteins under study. Essentially, the project has involved each of us studying the expression and localisation of a separate protein during the life cycle of Plasmodium, the malaria parasite. In addition, we aimed to study the morphology of the parasite when the protein was absent using a PTD construct. This removed expression of the protein from every stage in the life cycle, except for the blood stage where it is known to be essential. The point of studying the proteins was to identify a potential drug target for malaria treatment. The first thing we did was to transfect Plasmodium with a fusion gene, which coded for our protein of interest followed by the green fluorescence protein (GFP) tag. This would later allow us to see where our protein was in the cell. However, first we had to battle through the process of producing a Western blot, in order to show that the protein and GFP had been successfully fused. As Carrie has written about previously, this resulted in lots of failed attempts, with each one inevitably falling foul of a mistake that would leave us scratching our heads.

It's in there somewhere.

Life cycle imaging

Eventually though, we all had a band that we could confidently call our protein, and could then focus on the fluorescence imaging of the various Plasmodium life cycle stages. Because each attempt to transfect our PTDs into Plasmodium failed, this meant we could only study the protein localisation and not the phenotypes of the knockout. What I would consider to be the major difficulty in this part of the project was the time taken to culture the different life cycle stages. The process of growing Plasmodium in mouse blood takes around 3-4 days, after which time the mouse must be euthanised to prevent unnecessary suffering. The blood can then be used to image the parasite blood-stages (rings and trophozoites), or cultured for a further 24 hours to produce either schizonts or a mix of gametocytes, zygotes and ookinetes.

Plasmodium stages that can be imaged from mouse blood

Furthermore, studying the mosquito stages of the life cycle takes a total of 21 days following the collection of mouse blood. The mice must first be treated to eliminate all parasites except the gametocyte stages, then the mosquitoes are permitted to feed on them. At this point, the gametocytes in the mosquito gut become activated and leave the red blood cells. You then need to wait 10 days before some of the mosquitoes can be checked for oocysts lining the midgut, and mature oocysts can be imaged on day 14. After another week, on day 21, it should be possible to image sporozoites in the mosquito salivary glands. This completes the stages of the life cycle that can be studied, because the host's liver-stage parasite infections cannot be studied without complex dissection that would also render the mouse useless for study of the other stages. Unfortunately, the day 10 check of the mosquito infection for my protein showed no oocysts, meaning that it may not be possible to complete the oocyst and sporozoite imaging. This has therefore been a potential waste of 10 days, and would need the use of more mice and mosquitoes if a second attempt is to be made.

The Fluorescence Microscope

The actual process of taking parasite fluorescent images starts by mixing infected blood with DAPI, a DNA-specific stain that highlights the nucleus blue. In addition, the activated female, zygote and ookinete stages can also be tagged in red. This works by using a red fluorophore conjugated to an antibody the specifically binds to a protein expressed on the surface of these cell types. Once the fluorophores have been added, the blood can be searched for the desired stages under the fluorescence microscope. Any cells of interest must then be measured for their blue and red fluorescence, as well as the green fluorescence emitted by the GFP-tagged protein.

Activated female gametocyte with its membrane, nucleus and protein highlighted.

Getting a good image was always more difficult than you'd think, given that we had to make do with a single microscope shared between 4+ people, who often all had parasites to image at the exact same time. As well as this, we had to get the cell concentration exactly right, or they would be too sparse to find anything of interest or too dense to get the cell on its own. After many mice though, we were each able to create a complete collage to show the localisation of our protein at each of the ring-to-ookinete stages. All that remains now is to finish the study of the mosquito stages, and the project will be completed.

All in all there's been a lot of frustration and disappointment, but with some perseverance we got good results in the end. We also gained a lot from being able to adapt and learn from the things that went wrong, which in the end is a very important skill for a lab scientist to have. 

My (Our) project - Cell division in Plasmodium Berghei

What am I doing?

I am investigating the role of an Aurora-like protein kinase, PbARK2, in cell division in Plasmodium berghei, the rodent model for Plasmodium falciparum.

Why am I doing this?

Plasmodium undergoes cell division in a very atypical way – while most eukaryotes undergo open mitosis, in which the nuclear membrane breaks down before sister chromatids are pulled apart, Plasmodium cells maintain their nuclear envelope at all times. This is called closed mitosis. In addition to closed mitosis, their nuclear division is asynchronous – in a multinucleate cell, each nucleus divides independently of the ones around it. This suggests that there is no global control of mitosis within a cell, and a recent study highlighted that CDC20, a protein essential for mitotic regulation in all eukaryotes is in fact dispensable in the blood stages of parasite development.

Given that Plasmodium uses a drastically different system for cell division, many of the proteins involved may be very different to those in humans, making them good drug targets as off-target effects will be limited. By understanding the underlying biology of the system, we can better make predictions and investments into drug development.

How am I doing this?

ARK2 has already been shown to be essential for blood stage development in P. berghei. The plan was therefore as follows: place ARK2 under the control of an alternative promoter that is active in the blood stage to escape the lethal phenotype, allowing phenotypic analysis at later stages of development. This was to be achieved using a PTD construct (promoter trap using double homologous recombination), generated using molecular cloning techniques in the lab, before transfecting it into wild-type parasites.

Schematic representation of the PTD construct for placing Ark2 under the control of ama1

To provide additional insight into the function of ARK2, a previously generated parasite line expressing an ARK2-GFP tag was to be used to visualise the cellular distribution of the protein and determine expression patterns throughout the life cycle. This would be complimented by qPCR data produced for each life cycle stage.

What’s the story so far?

Last semester started pretty well – there are four of us MSci students in the lab doing very similar projects with different proteins so we felt like we weren’t all on our own. The initial molecular cloning went well, with the direct supervision of Declan, our resident lab-God, knower-of-all-things, provider-of-reagents and all-round-science-boss (the lab technician). With the 3’ fragment successfully inserted into the vector, we were left to do the 5’ section ourselves – this is where things started to go wrong. I managed to get lucky and only need to do one step twice, but Jack had to repeat his bacterial transformation four or five times. However, after a slightly shaky semester we had our constructs completed, sequenced to check accuracy, and ready for transfection. Two weeks before the end of the semester we were poised to begin our transfections and get on with the meat of the project. December 5^th came around, everything went well, nothing could have gone wrong, we felt confident and excited until a week later, when nothing survived the drug selection step, meaning our transfections hadn't worked.

Merry Christmas!

Four weeks later we were back and ready to get back into it. We knew the protocol better this time, we had read more about it, we wanted it more this time. And then it didn’t work. Neither did the third attempt, but the positive control worked just fine. Slightly disheartened, we decided to move on with getting our Western blots to show our protein was correct in the GFP-tagged lines. Carrie has a fantastic post telling you just how well that went… While a few weeks went by desperately hunting for meaning in the Rorschach tests we were creating, we started to learn how to use the fluorescence microscope to visualise our proteins. Jack has a detailed post talking about some of the trials and tribulations we experienced with the microscope, particularly the bit where more than four people need to use one microscope to do time sensitive investigations. Somewhere along the line we went from “just taking practice images” to being expected to produce a full collage of every life cycle stage – I’m still not sure exactly when that was, but I found that this was something I could confidently do. In addition to regular life cycle stages, we discovered that my protein was only transiently expressed during ookinete development, meaning we needed to image timepoints in ookinete development. This is where it gets exciting, because I got my first actual set of data usable in my write-up!

Collage of ookinete developmental stages showing punctate distribution of ARK2 dependent on stage.

Having finally got a decent set of images, I hoped that shortly after everything else would click into place – my Western would come out beautifully, my transfection would be successful and I’d have a clear picture of everything-ARK. However, the next transfection also failed and I still, 3 months and 14 Westerns later, don’t have a band for my protein. We’ve got a few plans for how to fix this but time is running out and it’s getting a bit worrying.

That being said, having spoken to my friends who are doing PhDs, and the post-docs in the lab, in research you have to get used to failure. The trick to being a good scientist is to persevere despite things not going right and work out how to fix it. Use the scientific method and think logically about what you can change, and maybe, just maybe, you’ll get the result you want.

If you’ve had a project come right down to the wire, or want to share any of your science woes please let me know in the comments so I can feel like it’s not just me that’s had this experience.

Taming your Western

During our projects we’ve learnt a multitude of molecular techniques, but the most elusive so far has been mastering the Western blot. Before Christmas we’d completed our promoter trap constructions, using molecular cloning to transfer our genes of interest (Kinesin13, Ark2, Rap9 and Smc3) under the control of the AMA1 promoter to restrict expression. We're currently waiting for our malaria transfections to show any sign of success, and for the meantime working on Western blotting and fluorescence imaging of our GFP-tagged proteins. On Ching has previously posted a summary of her project, so to avoid repeating ourselves I'd like to share one of the challenges of our project: the Western blot.  

The humble Western blot has many uses throughout biomedical research, including disease diagnosis, antibiotic efficacy studies, and the FIFA 2014 World Cup doping tests. For our purposes, we'd like to demonstrate that the fluorescence shown on our images indicates our protein of interest. Jack will shortly be writing a post about our experiences with microscopy, and I'd like to talk through the many mistakes we've made with Western blotting.

Image courtesy of David Taylor

In striving for the good blot, our initial results all too often resembled the bad and the ugly, and frankly things didn’t improve until we altered several steps of our lab protocol. We hope sharing our experiences troubleshooting Western Blotting will save some frustration for other students, and at the very least provide a snapshot of numerous ways it can go wrong. For any newcomers to Western Blotting - or anyone who just feels like punishing themselves -  we’d recommend this starter paper as a background to protein blotting.

1) Protein Handling – Lysis, Denaturation and Loading

The ugly: poor lysis, degradation
and overloading. 

Shown is one of our first gels, and to this day I still remember the look on our PIs face. The underlying nature of a Western relies on the quality of the protein that’ll be used. Our projects have involved isolating parasite proteins from infected mice blood, using CF11 columns for leukocyte depletion and then lysing cells to gain access to our proteins. During our first few westerns, we were having difficulty with protein degradation during cell lysis, which was compounded by chronic overloading of protein onto the gel. After several smeared and blotchy attempts, we rewrote the lab protocols for the lysis steps, lowering temperatures by lysing on ice and centrifuging at 4^oC, incorporating protease inhibitors and also restricting repeated freeze-thaw cycles.

Throughout this process we’ve also learnt to select detergents depending on our protein’s hypothesised cellular location – NP40 may be fine for cytoplasmic Kinesin 13, but for the nuclear Ark2 Sarkosyl would have been a better choice. A general rule is that the more compartments between you and your protein, the harsher the detergent needed. 

2) Supervising your Western

The overheated: a cautionary tale 

Borrowing the words of the Upturned Microscope: ”Your western blot hates you. Never leave it unsupervised”. After setting gels running, it’s usual to notice rising bubbles alongside fluctuating voltage and amp values. We’ve also found it’s a good idea to keep half an eye on our westerns, checking at least halfway through the cycle and not leaving it unsupervised for too long. During one of our earlier gels, there was an incident with overused electrophoresis buffer that resulted in the gel just melting within the chamber. A cursory glance over the tank will quickly reveal if everything is running as normal: checking the dye front is symmetrical, the tank feels cool to the touch and your protein hasn’t run off the gel edge will save a headache later.

The disturbed: temperature and
electrolyte fluctuations

If running for an extended period of time, it’s also best not to disturb the tank. For one of our experiments we were attempting to resolve a high molecular weight protein which refused to leave its well. We were running two gels, the first for 1hr 30min and the second for 2hrs, to reach better resolution. We made a rookie error: running the two in the same tank. Removing the first gel at the end of it's run meant turning off electrophoresis, removing the gel and refilling the electrolyte solution - all resulting in salt redistribution and temperature changes. The difference between the ladder on our two gels can clearly be seen – one resembles bands and the other blotchy smears.

3) Transfer

We’ve found that thoroughly removing any trapped air from the sandwich is an essential step once it’s been assembled onto the transfer system. Murphy ’s Law states that any bubbles occurring will appear just at the right height to obscure your protein band, and if you’re really unlucky these bubbles can span several lanes. Remembering to equilibrate sandwich components in transfer buffer for 2 – 10 minutes before assembling and then rolling out any air will help. This equilibration period also gives the gel time to cool which avoids transfer problems, and most buffers contain methanol to reduce air retention within your sandwich.

4) The Antibodies

Have you checked your primary antibody species matches your secondary? If you’ve ever had a western with no signal, this could be the culprit. Whilst this may seem like an obtuse mistake, it’s particularly easy to make when using Western Reagent kits. Our lab uses WesternBreeze by ThermoFisherScientific, which contains two delightfully similar bottles of mouse and rabbit antibodies, and once produced a confusingly blank blot. It is also possible to strip membranes and begin again from the beginning, but as this isn't something we've tried yet we'd love to hear our viewers comments on the process.

5) Background

The obscured: high background.

After our initial problems with high background on our westerns, we increased the frequency of our washing steps.If there’s too much background on the western, we’veincreasing washing steps will help to resolve the issue. However, if the background is disguising smearing or low signal to background levels, you may wish to examine how you’re handling the membrane and check your antibody dilutions. If assembling your sandwich involves the membrane sliding over the gel repeatedly (or when developing has frequent repositioning of your film within the cassette) it’s also possible to create smears and blotches.

If you’re having similar troubles with your western, we’d love to hear about it! You can share your worst westerns in the comments. For any students looking for a more comprehensive troubleshooting gallery, we’ve found the SDS-PAGE Hall of Shame curated by Rice University to be particularly inspiring. Join us again next fortnight when Jack shares his experiences with fluorescence microscopy in our next post: “Fluorescence and Failure”. 

References

1.  Baume N, Jan N, Emery C, et al (2015) 
   Antidoping programme and biological monitoring before and during the 2014 FIFA World Cup Brazil. British Journal of Sports Medicine. 49: 614-622
   
   
2. Kurien, B. T., & Hal Scofield, R. (2015). Western blotting. Methods. 38(4): 283–293 https://doi.org/10.1007/978-1-4939-2694-7
   

   
   
3.  Johnson, M. (2013). Detergents: Triton X-100, Tween-20, and More. Materials and Methods, [online] 3 (163). Available at: www.labome.com/method/Detergents-Triton-X-100-Tween-20-and-More.html.

How bacteria might win the fight against malaria, dengue and Zika

Have you heard of the most prevalent parasitic microbe in the world? 

Capable of infecting up to 70% of all insects? 

With a preference for murdering males in cold blood? 

Known as “Wolbachia”, this genus of bacteria is found in many arthropods, including those responsible for the transmission of many human pathogens; such as malaria, dengue fever, and the Zika virus. It gets transmitted vertically, meaning that infected females pass it directly to their offspring. Different strains of Wolbachia infect different hosts and can have many different effects. In filiarial nematodes Wolbachia plays a mutualistic role, with elimination of the symbiotic bacteria causing host sterility and sometimes death. In mosquitos however, Wolbachia has a more sinister role - causing something known as “cytoplasmic incompatibility”. I’ll explain what exactly that means in a minute, but the end result is that infected females will always produce infected offspring, and uninfected females have half the chance of producing offspring. This is a case where it’s easier to show, than tell.

This picture, taken from the Werren lab Wolbachia biology page, shows how cytoplasmic incompatibility leads to an increase in the proportion of infected hosts.

The exact mechanism for this cytoplasmic incompatibility (CI) isn’t yet clear. What is known is that some kind of modification must take place in the infected male during spermatogenesis, as mature sperm cells do not contain Wolbachia. We also know that rescue must occur at some point in the fertilised infected egg, as the presence of Wolbachia prevents CI from occurring, but uninfected eggs don’t produce offspring. The main consequence of CI is that the male pronucleus enters mitosis later than the female pronucleus, which means the genetic information from the male does not segregate properly during the first mitosis. This leads to the production of haploid daughter cells which is embryonically lethal, preventing any offspring from being formed.

In May last year, a paper by Daniel LePage et al., was published which found that just two genes in the Wolbachia genome were needed to cause CI. The sequences for these genes were found to originate from viral DNA, integrated from the bacteriophage WO genome at some distant point in Wolbachia’s evolutionary history. These two genes, named cifA and cifB (cytoplasmic incompatibility factors A and B), were initially identified as part of a pool of 113 genes shared between CI-causing strains of Wolbachia. This pool was narrowed down by removing proteins known to be very different or absent in a non-CI-causing strain, wAu, and by removing proteins known to be expressed by infected ovarian tissue as these might be generally expressed and not play a direct role in CI. This left just two genes, WD0631 and WD0632.

In section a, a Venn diagram shows 113 genes are shared between four common CI-causing strains of Wolbachia. In section b, a Venn diagram shows how these 113 genes were narrowed down to just two candidates for CI-causing genes – WD0631 and WD0632.

To investigate if these two were indeed the genes responsible for CI, transgenic lines of Drosophila melanogaster were created with both genes under the control of a promoter that is active in germ line cells. Males of this line were crossed with infected and uninfected females and the relative hatch rate calculated. It can be seen in the graph below that having both genes caused a significant change in the embryo hatch rate when crossed with uninfected females, and that this was rescued when crossed with infected females – as would be expected if these were the culprits for CI.

A comparison of relative embryo hatch rate in D. melanogaster. White symbols indicate uninfected males and females, black symbols indicate those infected with a CI-causing strain of Wolbachia.

Further to this the team investigated whether the cytological appearance was the same between Wolbachia infected embryos and transgenic WD0631⁺/WD0632⁺ lines, further strengthening their case that these were the responsible genes.

Panels a-f show different classifications of cytological appearance in recently fertilised ovaries of D. melanogaster. a) indicates unfertilised eggs, b) shows normal nucleated cells at 1h of development, c) shows normal embryos at 2h of development, d) shows failure of nuclear division after two to three mitoses, e) shows chromatin bridging and f) shows regional mitotic failure. In section g) the relative abundance of each cytological appearance is shown in different crossing scenarios.

The paper doesn’t go on to speculate on or provide any investigation into the mechanism of action of these two genes, but sets the stage for further work into this system and tantalisingly closes with the statement 

“Finally, cifA and cifB are important for arthropod pest and vector control strategies, as they could be an alternative or adjunct to current Wolbachia-based efforts aimed at controlling agricultural pests or curbing arthropod-borne transmission of infectious diseases”

This brings me on to the conclusion of this post and refers all the way back to the title – how can bacteria be used to fight insect-vector based disease?

There are already programs in place that are attempting to use Wolbachia-based methods of eradicating the insect vector of the dengue fever virus, the Aedes aegypti mosquito. By introducing large numbers of infected males into an uninfected population, the population can be reduced without the use of pesticides. The work done by LePage could provide the means to supplement Wolbachia in insect systems or possibly induce CI in other systems. An alternative to herbicides, or maybe maintenance-free pest control – can you think of any other applications? Let me know in the comments.

Ectogenesis

By Jack Sewell

The concept of ectogenesis, or in vitro pregnancy, is an interesting one, with huge ethical and social implications. Essentially it involves the use of an “artificial womb” machine that would facilitate the development of a foetus, by replicating the processes that normally happen during a natural pregnancy. This includes the provision of maternal blood or a suitable substitute, which could supply oxygen and nutrients to the foetus, as well as remove waste materials. To achieve this, an artificial placenta would have to mediate the transfer of these substances to and from the foetal circulation. Research in this field began in the 1980s, when in 1989 a human embryo was implanted in ex vivo uterus for the first time¹, though this line of study was quickly halted due to ethical concerns. Further advancements have been made since then, but currently there is no technology which is capable of supporting foetal development from conception through to birth.

Taken from motherboard.vice.com.

There are two potential uses for ectogenesis technology. The first would be as an improvement on current technology for the incubation of premature babies, which would likely give a much greater survival rate by more closely imitating the final stage of gestation. At present, incubators are only effective for babies born after at least 24 weeks of gestation¹. The second use is as an alternative to natural pregnancy, in which the majority or entirety of the gestation could be completed within the machine. An advantage that this could have over a natural pregnancy is the ability to monitor the developing foetus more closely throughout the process, perhaps as though it were on life support. This may be useful if the unborn baby is known or suspected to have a condition that would benefit from close monitoring. It could also make prenatal diagnosis tests safer and more viable, as the current tests (amniocentesis and chorionic villus tests) pose a small risk to the foetus.

Unsurprisingly, this technology raises significant ethical issues relating to the role of the mother and the reliance on technology as a substitute for a natural human process. As with many other scientific controversies, just because science can do something doesn’t necessarily mean that it should. Therefore, we must ask who - if and when this technology becomes available - should be allowed to use it to facilitate an entirely ex vivo pregnancy. For instance, should it only be available to those who cannot carry a child naturally, such as women with damaged uteri or gay men? In these cases, it would remove the need to find a willing surrogate mother, which is currently their only option if they wish to have a biological child. On the other hand, if the technology were made available to everyone, one could imagine that a pregnant woman may be offered the choice to transfer the embryo into an artificial womb for the remainder of the gestation. One ethical issue that arises here is whether this could mean taking away the control that women have over their own body and their own pregnancy. It may even lead to a world where artificial gestation is medically advised or legally enforced, due to reasons such as the poor health of the unborn child or the unfitness of the mother to take care of it during the pregnancy. Therefore, as this technology develops, there are serious decisions that lawmakers must prepare to make concerning the ethics of its use. For now though, there is plenty of time as it is estimated that the necessary technology for a full ex vivo pregnancy will not exist for several decades.

References

1. Bulletti, C., Palagiano, A., Pace, C., Cerni, A., Borini, A. and de Ziegler, D. The artificial womb. Ann. NY Acad. Sci. 1221, 124-128 (2011).

Is RTS,S the malaria vaccine we need?

The World Health Organisation has announced their intention to run three pilot projects within Sub-Saharan Africa for the first malaria vaccine. Due to begin in 2017, the pilots will will answer remaining questioning regarding RTS,S/AS01, which is the result of an enormous 32 years of research from partnerships between GlaxoSmithKline, the PATH Malaria Vaccine Initiative (MVI) and Walter Reed Army Institute of Research. Composed of a viral Hepatitis B surface antigen-like particle, an adjuvant to improve immunogenicity and a section of Plasmodium falciparum circumsporozoite protein, RTS,S represents the only candidate to have completed Phase II trials. 

Despite this success, vaccine efficacy has been found to fall far short of the 75% benchmark. With protection levels reported to be controversially low, what can we really expect from RTS,S?

Development timeline of the RTS,S vaccine shows the path from 1987 to 2014. Since creation of this figure, the vaccine has received a positive opinion from the European Medicines Agency and African countries are applying to national regulatory authorities for vaccine approval. 

The WHO seeks to answer these questions, with the pilot programmes aiming to "evaluate the feasibility of delivering the required 4 doses of RTS,S; the impact of RTS,S on lives saved; and the safety of the vaccine in the context of routine use". The replicability of protection levels has also been queried, with researchers advising four doses to provide maximum efficacy. This in itself could be problematic without comprehensive healthcare infrastructure. Summaries of the available Phase III trail data by Gosling and von Seidlein (2016) found the addition of the fourth booster justified, as in 6 - 12 week infants vaccine efficency rose from 18% to 25%, and in children 5 - 17 months old this rose from 28.3% to 36.3%. 

If RTS,S isn't the answer, where are the other candidates for malaria vaccines? Jack has previously posted regarding the types of malaria vaccine, and the WHO estimates the 20 candidates currently at various stages of the pipeline are all 5 - 10 years behind RTS,S. One of these candidates - PfSPZ, Sanaria - is composed of radiation-attentuated, whole sporozoites and has recently been reported to deliver full protection up to 25 weeks after innoculation. A recent review by Cowman et al (2016) highlights that this candidate lacks the stability of RTS,S, requiring liquid nitrogen to maintain viability which will undoubtedly be a logistical issue. It will be interesting to follow PfSPZ through more extensive trials to see if those statistics hold, but it should also be noted that this vaccine was also using a 4 dose schedule. 

Summarising the benefits and drawbacks of RTS,S, what can we expect from this vaccine? Whilst RTS,S could be integrated into current malaria control schemes, it is easy to be disappointed by the low levels of protection granted. At the beginning of this post I underlined the sheer amount of effort and time funneled into vaccine development, and for it still to fall short is hugely discouraging. Whilst the more cynical among us might argue that RTS,S is no solution in our fight against malaria, I find it important to consider the severity of the disease in sub-saharan Africa. Within this context any new tools are welcome, especially should we wish to continue reducing malarial burden. The WHO's decision to further test RTS,S is prudent but should they not seek to implement vaccination programs after pilot programmes have finalised it would be interesting to see if RTS,S would survive. If nothing else, the public and academic support for malaria vaccines generated by RTS,S should ease the passage of future vaccine candidates.

References

1. Gosling, R., and von Seidlein, L. (2016). The Future of the RTS,S/AS01 Malaria Vaccine: An Alternative Development Plan. PLoS Medicine 13(4) pp. 312-315
   
2. Ishizuka, A.S., Lyke, K.E., DeZure, A., Berry, A.A., Richie, T.L., Mendoza, F.H., Enama, M.E., Gordon, I.J., Chang, L.-J., Sarwar, U.N., et al. (2016). Protection against malaria at 1 year and immune correlates following PfSPZ vaccination. Nature Medicine 6, pp. 614–623.
   
3. Cowman, A.F., Healer, J., Marapana, D., and Marsh, K. (2016). Malaria: Biology and Disease. Cell 167, pp.610–624.

“Malaria drugs fail for first time on patients in UK”

A recent news article by the BBC reported that four cases of malaria in the UK failed to respond to artemether-lumefantrine treatment, the current front-line drug combination recommended by the WHO. Drug resistance to older antimalarial drugs, like chloroquine, is already widespread in many countries with endemic malaria. It spreads faster in areas where treatments have been dispensed inappropriately, because this fosters selection for resistant strains, similarly to antibiotic resistance. WWARN (worldwide antimalarial resistance network) has an excellent series of interactive maps that you can use to explore the available data on antimalarial resistance.

Of particular concern is resistance to artemisinins – a group of drugs which are the key component in the artemisinin-combined therapies recommended by the WHO. These are well tolerated therapies with high effectiveness, assuming the full course of treatment is taken. However, in certain areas where healthcare has not been well managed, or access to drugs is available without appropriate medical advice, artemisinin resistance has spread rapidly and pervasively. This is shown well in the map produced by Ashley et al., which demonstrates the prevalence of a certain cause of resistance in South East Asia.

A map produced by Ashley et al., showing prevalence of PfKelch13-associated artemisinin resistance in S.E. Asia, and the limited spread of PfKelch13-independent resistance in Africa.

The exact mechanism of resistance to artemisinin is still contested, with several ideas proposed so far but no consensus has yet been achieved. Part of the reason for this is that we still don’t really know how artemisinin kills the parasite in the first place… What we do know however, by using Genome Wide Association Studies of parasite genomes, is that mutations in the “Kelch-propeller domain” of Plasmodium falciparum Kelch13 are strongly associated with cases of artemisinin resistance. Kelch-propeller domains are used for protein:protein interactions, and can be very specific. K13 belongs to a superfamily of proteins known to mediate ubiquitin-regulated protein degradation and oxidative stress-responses, which plays into some theories suggesting that artemisinin acts by generating reactive oxygen species, thereby causing oxidative stress.

While mutations in PfK13 are certainly common in resistant cases (again, particularly in S.E. Asia), they aren’t constant. As seen in the map above, African cases of artemisinin-resistance are mostly not found to have any mutations in PfK13. Instead, variable artemisinin efficacy appears to be linked to multi-locus genotypes involving other resistance-associated proteins such as PfMDR1 (a multi-drug resistance pump) and PfCRT (a chloroquine resistance-associated transporter protein), along with less well understood proteins such as PfUBP1 and PfAP2MU. Worryingly, the diverse causes of artemisinin resistance suggest multiple, independent evolutions of resistance in isolated Plasmodium populations.

Going back to the original point of this post, the BBC article “Malaria drugs fail for the first time on patients in the UK”, we should look at the source of this information. This is a brief report of four case histories of patients with imported malaria that were initially treated with artemether-lumefantrine but presented with recurrent parasitaemia within 6 weeks of treatment with no intervening travel to malaria-endemic areas. In English, this means patients that caught malaria outside of the UK were treated to the point where they seemed healthy, but were then readmitted at a later date with the same symptoms without an opportunity to catch it again. Two patients had been to Uganda, one to Liberia and one to Angola – all in Africa. As you might be able to guess, when profiled for resistance markers, none of the parasites these patients were infected with had any PfKelch13 mutations. Instead, they had a variety of mutations in other loci known to be associated with resistance, shown in table 1.

Ultimately all patients were successfully treated with alternative drug combinations such as atovaquone/proguanil, quinine/doxycycline or artemether/lumefantrine with doxycycline so the initial headline may be slightly exaggerated when you take the full picture into account. However, this is still an important news story as it is the first time artemisinin resistance has been seen in the UK. As resistance spreads throughout Africa these cases will only become increasingly common, so finding appropriate alternatives is a necessary task. 

The BBC article includes an interview with Dr Sutherland, the lead researcher, who stresses that while this is not yet a national health crisis it is important for doctors to be aware that drugs may not work and says drug guidelines should be reviewed. He also suggested that large scale studies of drug efficacy need to be urgently undertaken in Africa to determine the severity and scale of the problem we could be facing very soon.

Personally, I feel that while this is a very important news story, the fact that just four UK cases made headlines while thousands occur every year abroad is telling of people’s attitudes in the UK. The spread of drug resistant malaria is very real and a very serious concern to people living in malaria endemic areas, causing death on a huge scale, but is not on the radar for most people until it directly concerns them. I think awareness needs to be raised about this as an issue that faces other countries, not just ours. I’d like to know if you agree or disagree, and if you have any ideas about ways we could achieve this goal – let me know in the comments.