This work was supported by the National Institutes of Health grant AI168307 to SPK.&#160; We thank the UGA CTEGD Flow Cytometry Core and the UGA CTEGD Microscopy Core. We also acknowledge the contributions of Ash Pathak, Anne Elliot, and the staff of UGA Sporocore in optimizing the protocol. We want to thank Dr. Daichi Kamiyama for valuable insights, discussion, and the parent plasmids containing GFP11 and GFP1-10. We would also like to thank members of the Kurup lab for their constant support, patience, and encouragement.

All research involving vertebrate animals in our laboratory was performed in compliance with the University of Georgia animal use guidelines and protocols.
1. Generation of P. berghei -infected mice
<ol>
	<li>Initiate blood-stage infection in male or female, 6-8-week-old C57BL/6 (B6) mice using wild-type P. berghei parasites. To do this, transfer cryopreserved P. berghei-infected blood (2 x 105 infected RBCs), reconstituted in 400 &#181;L of phosphate-buffered saline (PBS), into one or two donor mice intraperitoneally (i.p.).</li>
	<li>Transfer fresh blood collected through retro-orbital bleeding from the donor mice at 5 days post-infection (d.p.i.) into two na&#239;ve B6 mice. To do this, collect 200 &#181;L of blood, reconstitute with 300 &#181;L of PBS, and inject 200 &#181;L i.p. into the recipient mice. Ensure that the parasitemia in the donors at this time point is 2%-5%19,20.</li>
	<li>Monitor parasitemia as previously described19,20, starting from 2 d.p.i. in the recipients. When parasitemia ranges from 3%-5% (around 5 d.p.i.), euthanize the recipient mice and then collect blood through cardiac puncture or retro-orbital bleeding. Euthanize the mice using carbon dioxide inhalation followed by cervical dislocation, or according to institutional guidelines. 
	&#8203;NOTE: Once parasitemia exceeds 5%, the transfection efficiency may be reduced due to the increased prevalence of multiply infected RBCs.</li>
</ol>
2. Generation of schizonts in culture
<ol>
	<li>Collect blood from the infected mice in step 1.3 (approximately 2 mL in total from two mice) into 5 mL of Alsever&#39;s solution (see Table of Materials) in a sterile environment. Perform steps 2.1 to 3.12 under sterile conditions. 
	NOTE: Sterile conditions include wearing gloves, wiping gloves and surfaces with 70% ethanol, employing sterile consumables and materials, and working within a biosafety cabinet.</li>
	<li>Centrifuge the blood for 8 min at 450 x g at room temperature (RT). Discard the supernatant and resuspend the cell pellet in 10 mL of complete RPMI media (RPMI 1640 with 20% fetal bovine serum [FBS] and 1% penicillin-streptomycin, v/v).</li>
	<li>Centrifuge for 8 min at 450 x g at RT. Resuspend the pellet in 25 mL of complete RPMI media and transfer to a T75 culture flask with a filter cap.</li>
	<li>Place in an incubator at 36.5 &#176;C and 5% CO2, while gently shaking to keep the cells in suspension for 16 h.</li>
	<li>At the 16 h time point, check for schizont development. To do this, collect 500 &#181;L of the sample, centrifuge at 450 x g for 8 min, resuspend the pellet in 10 &#181;L of complete RPMI media, and prepare a thin blood smear. Stain the blood smear with Giemsa stain19 (see Table of Materials) and determine the frequency of schizonts (Figure 1).</li>
	<li>For optimal transfection efficiency, 60%-70% of the parasites should be in the schizont stage. If fewer than this threshold is determined, continue culturing as in step 2.4 and check every hour to ensure the above threshold is crossed before proceeding.</li>
</ol>
3. Transfection of Plasmodium schizonts
<ol>
	<li>Prepare a gradient centrifugation buffer by reconstituting 13 mL of density gradient stock medium (see Table of Materials) with 1.44 mL of 1.5 M NaCl and 5.6 mL of 0.15 M NaCl in a 50 mL centrifugation tube.</li>
	<li>Transfer 25 mL of blood culture to a fresh 50 mL centrifugation tube. Carefully layer 20 mL of gradient centrifugation buffer to the bottom of the centrifugation tube using a narrow glass pipette to create a gradient column. Take care not to disturb the liquid layers and interfaces and ensure a clear division between the buffer and the media.</li>
	<li>Centrifuge for 20 min at 450 x g with no brake at RT. Using a Pasteur pipette, carefully remove the schizont layer15 located at the interface of the culture media and the gradient centrifugation buffer, and place it in a new 50 mL centrifugation tube.</li>
	<li>Resuspend the isolated schizonts in complete RPMI media and bring up the total volume to 40 mL. Centrifuge at 450 x g for 8 min at RT and discard the supernatant.</li>
	<li>Resuspend the schizonts in 10 mL of complete RPMI media. Then, transfer 1 mL of this solution into a 1.5 mL microcentrifuge tube for each transfection, and centrifuge at 16,000 x g for 5 s to pellet the infected RBCs. The infected RBCs derived from two mice in the above manner can be used for up to 10 separate transfections.</li>
	<li>Combine 100 &#181;L of the electroporation buffer (nucleofector solution from the electroporation kit; see Table of Materials) at RT with 5 &#181;g of circular or linearized target plasmid DNA (in up to a maximum of 10 &#181;L volume), as applicable for each transfection in separate 1.5 mL microcentrifuge tubes. A &#34;no plasmid&#34; sample must be included as a control for transfection and antibiotic selection.</li>
	<li>Remove the supernatant after centrifugation from step 3.5 and carefully resuspend the infected RBCs in the prepared nucleofector solution + plasmid DNA (from step 3.6).</li>
	<li>Transfer the suspension (nucleofector solution + plasmid + schizonts; 110 &#181;L maximum) into electroporation cuvettes. Transfect using a suitable nucleofector program (U-033, see Table of Materials). 
	NOTE: Ensure that steps 3.8 to 3.10 are performed at RT.</li>
	<li>Add 100 &#181;L of complete RPMI media to the cuvette and transfer the entire solution (now 200 &#181;L total volume) into a 1.5 mL microcentrifuge tube and keep at RT.</li>
	<li>Inject the 200 &#181;L solution of transfected parasites into mice intravenously (i.v.), working to minimize the time between transfection and the final injection into mice. 
	&#8203;NOTE: Transfected schizonts are injected into mice less than 5 min after electroporation to maximize the efficiency of the protocol.</li>
</ol>
4. Selection of the transfected parasites
<ol>
	<li>Check the parasitemia levels in the mice inoculated with the transfected schizonts daily from 2 d.p.i. to verify infection.</li>
	<li>Once the infection is verified, start drug selection using oral administration of the drug in drinking water, offered ad libitum. 
	NOTE: Pyrimethamine was employed as the selectable drug marker for the plasmid. In this case, 17.5 mg of pyrimethamine was added to 250 mL of clean water (final concentration of 70 mg/L). In addition, 10 g of sugar was added to this water to encourage adequate consumption of the pyrimethamine-containing water by mice.</li>
	<li>Monitor parasitemia every two days using blood smears made with 5 &#181;l of blood, starting from 6 days after the initiation of pyrimethamine treatment. The lack of detectable parasites after 15 days indicates a failed transfection; in this case, reinitiate the process from step 1.1 for a fresh attempt.</li>
	<li>When parasitemia reaches at least 1%, transfer the parasites to na&#239;ve B6 mice for the creation of blood-stage parasite stocks. Continue selection in the newly infected mice until parasitemia reaches 2%-5%, at which point generate stocks and cryopreserve them21.</li>
</ol>
5. Infection of mosquitoes with the transgenic lines
<ol>
	<li>Infect B6 mice with 200 &#181;L of blood containing the selected parasites (2 x 105 infected RBCs/mouse, i.v.). Determine parasitemia in these mice on the day of mosquito feeding and infection. Parasitemia between 2%-5% is optimal for the infection of mosquitoes. Once verified, immediately transfer the infection to the female Anopheles stephensi mosquitoes according to steps 5.2-5.4). 
	NOTE: Incubators containing mosquitoes for infection should be maintained at 20.5&#160;&#176;C, at 80% humidity, and on a 12 h light/dark cycle (lights on between 07:00 a.m. to 07:00 p.m.).</li>
	<li>The day before mosquito infection, separate the female mosquitoes by placing a heat source on top of the cage containing both male and female mosquitoes, such as a thin plastic bladder or glove filled with water heated to 37 &#176;C. Remove the female mosquitoes, which are attracted to the heat source, using an insect aspirator and transfer these to a separate, smaller cage for infection. 
	NOTE: Male mosquitoes can be distinguished from female mosquitoes by their slightly reduced body size and plumose antennae.</li>
	<li>Inject the infected mice with a general anesthetic (e.g., 2% tribromoethanol/avertin). Ensure complete anesthesia by firmly pinching the foot pad of each mouse. If they do not react, they are sufficiently sedated and ready to transfer the infection into mosquitoes.</li>
	<li>Place the anesthetized mice on top of the caged female mosquitoes (from step 5.2) under sternal recumbency. Manually spread the front and back legs to provide the greatest surface area for mosquito feeding. Leave the infected mice over each cage for a total of 15 min, checking every 5 min to ensure the mice are not awake. Once feeding is complete, euthanize the mice using cervical dislocation, or as per the institutional animal use protocol, and dispose of them safely.</li>
</ol>
6. Collection of sporozoites
<ol>
	<li>Check mosquito midguts for oocysts at approximately 10 d.p.i. to verify successful infection and the development of Plasmodium within the mosquitoes. Isolate mosquito midguts from the abdomen by removing the terminal abdominal segment using forceps. Visualize the midguts under polarized light for the presence of oocysts22.</li>
	<li>Wild-type sporozoites are expected to be present in the salivary glands of mosquitoes 18-21 days after feeding on infected mice. At day 18, dissect 5-10 mosquitoes to assess sporozoite numbers. 
	NOTE: Mosquitos are washed and killed in ethanol. Subsequently, dead mosquitos are washed twice with PBS to remove debris prior to dissection.</li>
	<li>To isolate and count the sporozoites, carefully remove the head of the mosquito while applying pressure to the mosquito thorax using forceps or a fine dissecting needle. The salivary glands remain attached to the removed head (Figure 2).</li>
	<li>Remove any additional mosquito debris from the salivary glands and place in a microcentrifuge tube containing 400 &#181;L of mosquito dissection media (Dulbecco&#39;s modified Ealge medium [DMEM] with 1% mouse serum, 100 U/mL penicillin, and 100 &#181;g/mL streptomycin; see Table of Materials).</li>
	<li>Disrupt the isolated salivary glands by passing through a 30 G needled syringe 15-20 times. Place 10 &#181;L of the disrupted salivary glands in mosquito dissection media into a hemocytometer and count. Resuspend in the desired concentration of mosquito dissection media or PBS for injection into mice, inoculation into cell cultures, or long-term cryopreservation23,24.</li>
</ol>

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The authors declare no conflict of interest.

We have used the above protocol in our laboratory to create several lines of transgenic P. berghei parasites. Though optimized for P. berghei, we have also successfully used this protocol to generate transgenic P. yoelii sporozoites. After injecting the transfected schizonts into mice, parasites are detectable typically no later than 3 d.p.i. in all groups, including the no plasmid control. Selection is started only once parasitemia has been detected to ensure the viability of parasites followi...

Despite advances in drug development and research into malaria prevention and treatment, the global disease burden of malaria remains high. Over half a million people die of malaria each year, with the highest levels of mortality seen among children living in malaria-endemic regions, such as sub-Saharan Africa1. Malaria is caused by the parasite Plasmodium, which is transmitted to humans through the bite of female Anopheles mosquitoes bearing the parasite in their salivary glands. The infectious stage of Plasmodium-the sporozoites-are deposited in the skin of the vertebrate hosts during a blood meal and travel through the bloodstream to infect liver cells, where they undergo mandatory development (constituting pre-erythrocytic malaria) prior to infecting the erythrocytes. The infection of the erythrocytes initiates the blood-stage of malaria and is responsible for the entirety of the morbidity and mortality associated with the disease2,3.
The obligate nature of the pre-erythrocytic development of Plasmodium has made it an attractive target for prophylactic vaccine and drug development efforts4. A prerequisite for studying the biology of pre-erythrocytic malaria, as well as the development of vaccines or drugs targeting the liver stage, is access to Plasmodium sporozoites. Furthermore, our ability to generate genetically modified Plasmodium sporozoites has been instrumental in the success of such research endeavors5,6,7,8,9. Transgenic Plasmodium lines expressing fluorescent or luminescent reporter proteins have allowed us to track their development in vivo and in vitro10,11. Genetically attenuated parasites (GAPs), generated through the deletion of multiple genes in Plasmodium, are also some of the most promising vaccine candidates12,13.
Rodent and non-human primate malaria models have helped us understand the mechanisms of host-parasite interactions in human malaria due to the similarities in biology and life cycle among Plasmodium species14. The use of Plasmodium species that infect rodents, but not humans (e.g., P. berghei) allows the maintenance of the complete parasite life cycle and the generation of infectious sporozoites for studying liver-stage malaria in a controlled, biosafety level 1 setting. A variety of separate protocols already exist for the generation of transgenic blood-stage Plasmodium parasites15, infection of mosquitoes16, and isolation of sporozoites17. Here, we outline a comprehensive protocol combining these methodologies in order to generate and isolate transgenic P. berghei sporozoites, utilizing the novel transgenic strain PbGFP11 as an example.&#160;PbGFP11 traffics the 11th &#946;-strand of super-folder green fluorescent protein (GFP), GFP11,&#160;into the parasitophorous vacuole (PV) generated in the host hepatocytes. PbGFP11&#160;is used in conjunction with transgenic hepatocytes (Hepa1-6 background) expressing residues that constitute the GFP 1-10 fragment (GFP1-10) in the cytoplasm (Hepa GFP1-10 cells). PbGFP11 reports PV lysis in the host hepatocytes through self-complementation and the reformation of functional GFP and the green fluorescence signal18. Of note, GFP11 is encoded as a series of seven tandem sequences in PbGFP11&#160;to enhance the resulting fluorescence signal. Upon staining PbGFP11&#160;sporozoites with the cytoplasmic dye CellTrace Violet (CTV), we can track the parasites. The lysis of such CTV-stained intracellular parasites itself results in leakage of CTV into the host cell cytoplasm and staining of the host cell. In addition to visualizing and distinguishing the lysis of Plasmodium PV and/or the parasite in host hepatocytes, this system can be reliably used to study the immune pathways responsible for either of these processes, through the genetic or therapeutic perturbation of the molecular components of such pathways.

<table><tbody><tr><td>30 G x 1/2&quot; Syringe needle</td><td>Exel international</td><td>26437</td><td></td></tr><tr><td>Alsever&#039;s solution</td><td>Sigma-Aldritch</td><td>A3551-500ML</td><td></td></tr><tr><td>Amaxa Basic Parasite Nucleofector Kit 2</td><td>Lonza</td><td>VMI-1021</td><td></td></tr><tr><td>Avertin (2,2,2-Tribromoethanol)</td><td>TCI America</td><td>T1420</td><td></td></tr><tr><td>Blood collection tubes</td><td>BD bioscience</td><td>365967</td><td>for serum collection</td></tr><tr><td>C-Chip disposable hematocytometer</td><td>INCYTO</td><td>DHC-N01-5</td><td></td></tr><tr><td>CellVeiw Cell Culture Dish</td><td>Greiner Bio-One</td><td>627860</td><td></td></tr><tr><td>Centrifuge 5425</td><td>Eppendorf</td><td>5405000107</td><td></td></tr><tr><td>Centrifuge 5910R</td><td>Eppendorf</td><td>5910R</td><td>For gradient centrifugation</td></tr><tr><td>Delta Vision II - Inverted microscope system</td><td>Olympus</td><td>IX-71</td><td></td></tr><tr><td>Dimethyl Sulfoxide</td><td>Sigma</td><td>D5879-500ml</td><td></td></tr><tr><td>Fetal bovine serum</td><td>GenClone</td><td>25-525</td><td></td></tr><tr><td>GFP11 plasmid</td><td>Kurup Lab</td><td>pSKspGFP11</td><td>Generated from PL0017 plasmid</td></tr><tr><td>Giemsa Stain</td><td>Sigma-Aldritch</td><td>48900-1L-F</td><td></td></tr><tr><td>Hepa GFP1-10 cells</td><td>Kurup Lab</td><td>Hepa GFP1-10</td><td>Generated from Hepa 1-6 cells (ATCC Cat# CRL-1830)</td></tr><tr><td>Mouse Serum</td><td></td><td></td><td>Used for mosquito dissection media</td></tr><tr><td>NaCl</td><td>Millipore-Sigma</td><td>SX0420-5</td><td>1.5 M and 0.15 M for percoll solution</td></tr><tr><td>Nucleofector II</td><td>Amaxa Biosystems (Lonza)</td><td></td><td>Program U-033 used for RBC electroporation</td></tr><tr><td>Pasteur pipette</td><td>VWR</td><td>14673-043</td><td></td></tr><tr><td>Penicillin/Streptomycin</td><td>Sigma-Aldritch</td><td>P0781-100ML</td><td></td></tr><tr><td>Percoll (Density gradient stock medium)</td><td>Cytivia</td><td>17-0891-02</td><td>Details in protocol</td></tr><tr><td>PL0017 Plasmid</td><td>BEI Resources</td><td>MRA-786</td><td></td></tr><tr><td>Pyrimethamine (for oral administration)</td><td>Sigma</td><td>46706</td><td>Preparation details: Add 17.5 mg Pyrimethamine to 2.5 mL of DMSO. Vortex, if needed to dissolve completely; Adjust pH of 225 mL of dH&lt;sub&gt;2&lt;/sub&gt;O to 4 using HCL. Add Pyrimethamine in DMSO to water and bring to 250 mL. Add 10 g of sugar to encourage regular consumption of drugged water. Pyrimethamine is light sensitive. Use dark bottle or aluminum foil covered bottle when treating mice.</td></tr><tr><td>RPMI 1640</td><td>Corning</td><td>15-040-CV</td><td></td></tr><tr><td>SoftWoRx microscopy software</td><td>Applied Precision</td><td>v6.1.3</td><td></td></tr></tbody></table>

generating genetically modified plasmodium berghei sporozoites

Malaria is a deadly disease caused by the parasite Plasmodium and is transmitted through the bite of female Anopheles mosquitoes. The sporozoite stage of Plasmodium deposited by mosquitoes in the skin of vertebrate hosts undergoes a phase of mandatory development in the liver before initiating clinical malaria. We know little about the biology of Plasmodium development in the liver; access to the sporozoite stage and the ability to genetically modify such sporozoites are critical tools for studying the nature of Plasmodium infection and the resulting immune response in the liver. Here, we present a comprehensive protocol for the generation of transgenic Plasmodium berghei sporozoites. We genetically modify blood-stage P. berghei and use this form to infect Anopheles mosquitoes when they take a blood meal. After the transgenic parasites undergo development in the mosquitoes, we isolate the sporozoite stage of the parasite from the mosquito salivary glands for in vivo and in vitro experimentation. We demonstrate the validity of the protocol by generating sporozoites of a novel strain of P. berghei expressing the green fluorescent protein (GFP) subunit 11 (GFP11), and show how it could be used to investigate the biology of liver-stage malaria.

Determining the frequency and development of schizonts is critical for assuring that enough viable parasites are in the optimal stage for transfection. Immature schizonts can be differentiated from fully mature schizonts by the presence of fewer merozoites that do not fill the entire intracellular space of the RBC (Figure 1B). It is important to note that when making blood smears from cultured blood, infected RBCs may break open, resulting in the observation of free, extracellular merozoites...

Watch this Scientific Journal Video about Generating Genetically Modified Plasmodium berghei Sporozoites at JoVE.com

Generating Genetically Modified Plasmodium berghei Sporozoites

Malaria is transmitted through inoculation of the sporozoite stage of Plasmodium by infected mosquitoes. Transgenic Plasmodium has allowed us to understand the biology of malaria better and has contributed directly to malaria vaccine development efforts. Here, we describe a streamlined methodology to generate transgenic Plasmodium berghei sporozoites.

Generating Genetically Modified Plasmodium berghei Sporozoites

Malaria is a deadly disease caused by the parasite Plasmodium and is transmitted through the bite of female Anopheles mosquitoes. The sporozoite stage of ...

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Research

JoVE Journal

Immunology and Infection

1.1K Views.  University of Georgia. Malaria is transmitted through inoculation of the sporozoite stage of Plasmodium by infected mosquitoes. Transgenic Plasmodium has allowed us to understand the biology of malaria better and has contributed directly to malaria vaccine development efforts. Here, we describe a streamlined methodology to generate transgenic Plasmodium berghei sporozoites.

Video: Generating Genetically Modified Plasmodium berghei Sporozoites

Protocol for Production of a Genetic Cross of the Rodent Malaria Parasites

Variation in response to antimalarial drugs and in pathogenicity of malaria parasites is of biologic and medical importance. Linkage mapping has led to successful identification of genes or loci underlying various traits in malaria parasites of rodents1-3 and humans4-6. The malaria parasite Plasmodium yoelii is one of many malaria species isolated from wild African rodents and has been adapted to grow in laboratories. This species reproduces many of the biologic characteristics of the human malaria parasites; genetic markers such as microsatellite and amplified fragment length polymorphism (AFLP) markers have also been developed for the parasite7-9. Thus, genetic studies in rodent malaria parasites can be performed to complement research on Plasmodium falciparum. Here, we demonstrate the techniques for producing a genetic cross in P. yoelii that were first pioneered by Drs. David Walliker, Richard Carter, and colleagues at the University of Edinburgh10.

Genetic crosses in P. yoelii and other rodent malaria parasites are conducted by infecting mice Mus musculus with an inoculum containing gametocytes of two genetically distinct clones that differ in phenotypes of interest and by allowing mosquitoes to feed on the infected mice 4 days after infection. The presence of male and female gametocytes in the mouse blood is microscopically confirmed before feeding. Within 48 hrs after feeding, in the midgut of the mosquito, the haploid gametocytes differentiate into male and female gametes, fertilize, and form a diploid zygote (Fig. 1). During development of a zygote into an ookinete, meiosis appears to occur11. If the zygote is derived through cross-fertilization between gametes of the two genetically distinct parasites, genetic exchanges (chromosomal reassortment and cross-overs between the non-sister chromatids of a pair of homologous chromosomes; Fig. 2) may occur, resulting in recombination of genetic material at homologous loci. Each zygote undergoes two successive nuclear divisions, leading to four haploid nuclei. An ookinete further develops into an oocyst. Once the oocyst matures, thousands of sporozoites (the progeny of the cross) are formed and released into mosquito hemoceal. Sporozoites are harvested from the salivary glands and injected into a new murine host, where pre-erythrocytic and erythrocytic stage development takes place. Erythrocytic forms are cloned and classified with regard to the characters distinguishing the parental lines prior to genetic linkage mapping. Control infections of individual parental clones are performed in the same way as the production of a genetic cross.

Genetic crosses of rodent malaria parasites are performed by feeding two genetically distinct parasites to mosquitoes. Recombinant progeny are cloned from mouse blood after allowing mosquitoes to bite infected mice. This video shows how to produce genetic crosses of Plasmodium yoelii and is applicable to other rodent malaria parasites.

Variation in response to antimalarial drugs and in pathogenicity of malaria parasites is of biologic and medical importance. Linkage mapping has led to ...

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

Malaria is a significant cause of morbidity and mortality worldwide. This disease, which primarily affects those living in tropical and subtropical regions, is caused by infection with Plasmodium parasites. The development of more effective drugs to combat malaria can be accelerated by improving our understanding of the biology of this complex parasite. Genetic manipulation of these parasites is key to understanding their biology; however, historically the genome of P. falciparum has been difficult to manipulate. Recently, CRISPR/Cas9 genome editing has been utilized in malaria parasites, allowing for easier protein tagging, generation of conditional protein knockdowns, and deletion of genes. CRISPR/Cas9 genome editing has proven to be a powerful tool for advancing the field of malaria research. Here, we describe a CRISPR/Cas9 method for generating glmS-based conditional knockdown mutants in P. falciparum. This method is highly adaptable to other types of genetic manipulations, including protein tagging and gene knockouts.

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

We describe a method for generating glmS-based conditional knockdown mutants in Plasmodium falciparum using CRISPR/Cas9 genome editing.

Malaria is a significant cause of morbidity and mortality worldwide. This disease, which primarily affects those living in tropical and subtropical ...

Genetics

Myeloid Cell Isolation from Mouse Skin and Draining Lymph Node Following Intradermal Immunization with Live Attenuated Plasmodium Sporozoites

Malaria infection begins when the sporozoite stage of Plasmodium is inoculated into the skin of a mammalian host through a mosquito bite. The highly motile parasite not only reaches the liver to invade hepatocytes and transform into erythrocyte-infective form. It also migrates into the skin and to the proximal lymph node draining the injection site, where it can be recognized and degraded by resident and/or recruited myeloid cells. Intravital imaging reported the early recruitment of brightly fluorescent Lys-GFP positive leukocytes in the skin and the interactions between sporozoites and CD11c+ cells in the draining lymph node. We present here an efficient procedure to recover, identify and enumerate the myeloid cell subsets that are recruited to the mouse skin and draining lymph node following intradermal injection of immunizing doses of sporozoites in a murine model. Phenotypic characterization using multi-parametric flow cytometry provides a reliable assay to assess early dynamic cellular changes during inflammatory response to Plasmodium infection.

Myeloid Cell Isolation from Mouse Skin and Draining Lymph Node Following Intradermal Immunization with Live Attenuated Plasmodium Sporozoites

We describe here a protocol for isolating myeloid cells from mouse skin and draining lymph node following intradermal injection of Plasmodium sporozoites. Flow cytometry of collected cells provides a reliable assay to characterize the skin and draining lymph node inflammatory response to the parasite.

Malaria infection begins when the sporozoite stage of Plasmodium is inoculated into the skin of a mammalian host through a mosquito bite. The highly ...

Phenotypic Analysis of Rodent Malaria Parasite Asexual and Sexual Blood Stages and Mosquito Stages

Recent advances in genetics and systems biology technologies have promoted our understanding of the biology of malaria parasites on the molecular level. However, effective malaria parasite targets for vaccine and chemotherapy development are still limited. This is largely due to the unavailability of relevant and practical in vivo infection models for human Plasmodium species, most notably for P.&#160;falciparum and P. vivax. Therefore, rodent malaria species have been extensively used as practical alternative in vivo models for malaria vaccine, drug targeting, immune response, and functional characterization studies of conserved Plasmodiumspp. genes. Indeed, rodent malaria models have proven to be invaluable, especially for exploring mosquito transmission and liver stage biology, and were indispensable for immunological studies. However, there are discrepancies in the methods used to evaluate the phenotypes of transgenic and wild-type asexual and sexual blood-stage parasites.&#160;Examples of these discrepancies are the choice of an intravenous vs. intraperitoneal infection of rodents with blood-stage parasites and the evaluation of male gamete exflagellation. Herein, we detail standardized experimental methods to evaluate the phenotypes of asexual and sexual blood stages in transgenic parasites expressing reporter-gene or wild-type rodent malaria parasite species. We also detail the methods to evaluate the phenotypes of malaria parasite mosquito stages (gametes, ookinetes, oocysts, and sporozoites) inside Anopheles mosquito vectors. These methods are detailed and simplified here for the lethal and non-lethal strains of P. berghei and P. yoelii but can also be applied with some adjustments to P. chabaudi and P. vinckei rodent malaria species.

Due to the striking similarities of the life cycle and biology of rodent malaria parasites to human malaria parasites, rodent malaria models have become indispensable for malaria research. Herein, we standardized some of the most important techniques used in the phenotypic analysis of wild-type and transgenic rodent malaria species.

Recent advances in genetics and systems biology technologies have promoted our understanding of the biology of malaria parasites on the molecular level. ...

In Vivo Tracking of Edema Development and Microvascular Pathology in a Model of Experimental Cerebral Malaria Using Magnetic Resonance Imaging

Cerebral malaria is a sign of severe malarial disease and is often a harbinger of death. While aggressive management can be life-saving, the detection of cerebral malaria can be difficult. We present an experimental mouse model of cerebral malaria that shares multiple features of the human disease, including edema and microvascular pathology. Using magnetic resonance imaging (MRI), we can detect and track the blood-brain barrier disruption, edema development, and subsequent brain swelling. We describe multiple MRI techniques that can visualize these pertinent pathological changes. Thus, we show that MRI represents a valuable tool to visualize and track pathological changes, such as edema, brain swelling, and microvascular pathology, in vivo.

In Vivo Tracking of Edema Development and Microvascular Pathology in a Model of Experimental Cerebral Malaria Using Magnetic Resonance Imaging

We describe a mouse model of experimental cerebral malaria and show how inflammatory and microvascular pathology can be tracked in vivo using magnetic resonance imaging.

Cerebral malaria is a sign of severe malarial disease and is often a harbinger of death. While aggressive management can be life-saving, the detection of ...

Medicine

Plasmodium falciparum Gametocyte Culture and Mosquito Infection Through Artificial Membrane Feeding

Malaria remains one of the most important public health problems, causing significant morbidity and mortality. Malaria is a mosquito borne disease transmitted through an infectious bite from the female Anopheles mosquito. Malaria control will eventually rely on a multitude of approaches, which includes ways to block transmission to, through and from mosquitoes. To study mosquito stages of malaria parasites in the laboratory, we have optimized a protocol to culture highly infectious Plasmodium falciparum gametocytes, a parasite stage required for transmission from the human host to the mosquito vector. P. falciparum gametocytes mature through five morphologically distinct steps, which takes approximately 1-2 weeks. Gametocyte culture described in this protocol is completed in 15 days and are infectious to mosquitoes from days 15-18. These protocols were developed to maintain a continuous cycle of infection competent gametocytes and to maintain uninterrupted supply of mosquito stages of the parasite. Here, we describe the methodology of gametocyte culture and how to infect mosquitoes with these parasites using glass membrane feeders.

Plasmodium falciparum Gametocyte Culture and Mosquito Infection Through Artificial Membrane Feeding

Detailed investigations on mosquito stages of malaria parasites are critical to design effective transmission blocking strategies. This protocol demonstrates how to effectively culture infectious gametocytes and then feed these gametocytes to mosquitoes to generate mosquito stages of P. falciparum.

Malaria remains one of the most important public health problems, causing significant morbidity and mortality. Malaria is a mosquito borne disease ...

Biology

Detection of Plasmodium Sporozoites in Anopheles Mosquitoes using an Enzyme-linked Immunosorbent Assay

Plasmodium sporozoites are the infective stage of malaria parasites that infect humans. The sporozoites residing in the salivary glands of female Anopheles mosquitoes are transmitted to humans via mosquito bites during blood feeding. The presence of sporozoites in the mosquito salivary glands thus defines mosquito infectiousness. To determine whether an Anopheles mosquito carries Plasmodium sporozoites, the enzyme-linked immunosorbent assay (ELISA) method has been the standard tool to detect the Plasmodium circumsporozoite protein (CSP), the major surface protein of the sporozoites. In this method, the head along with the thorax of each mosquito is separated from the abdomen, homogenized, and subjected to a sandwich ELISA to detect the presence of CSP specific to Plasmodium falciparum and each of the two subtypes, VK210 and VK247, of Plasmodium vivax.This method has been used to study malaria transmission, including the seasonal dynamics of mosquito infection and the species of the major malaria vectors in the study sites.

Detection of Plasmodium Sporozoites in Anopheles Mosquitoes using an Enzyme-linked Immunosorbent Assay

This protocol describes a sandwich enzyme-linked immunosorbent assay to detect salivary gland sporozoites in mosquitoes. Using easily available monoclonal antibodies, the method enables cost-effective, high-throughput detection of mosquitoes carrying Plasmodium falciparum or Plasmodium vivax. The method is suitable for malaria transmission research, including vector surveys.

Plasmodium sporozoites are the infective stage of malaria parasites that infect humans. The sporozoites residing in the salivary glands of female ...

Identifying Kinases Rewiring in Malaria Parasite: A Chemical Genetics Approach

Understanding the Development of Compensatory Pathways in a Mutant Malaria Parasite Harbouring Hypomorphic Allele of Plant-Like Kinases

One of the mechanisms for subverting the effect of drugs by&#160;the malaria parasite is through rewiring of its transcriptome. The effect is more pronounced for target genes belonging to the multigene family. Plasmodium falciparum protein kinases belonging to the CDPK family are essential for blood stage development. As such, CDPKs are considered good targets for the development of anti-malarial compounds. The chemical genetics approach has been historically used to elucidate the function of protein kinases in higher eukaryotes. It requires the substitution of gatekeeper residue for another amino acid with a different side chain through genetic manipulation. Amino acid substitution at the gatekeeper position modulates the activity of a protein kinase and changes its susceptibility to a specific class of compounds known as bumped kinase inhibitors (BKIs) that help in the functional identification of the target gene. Here, we have exploited the chemical genetics approach to understand compensatory mechanisms evolved by a mutant parasite harboring a hypomorphic allele of cdpk1. Overall, our approach helps in identifying compensatory pathways that may be simultaneously targeted to prevent the development of drug resistance against individual kinases.

Author Spotlight: Identifying Compensatory Pathways in Malaria Parasites Containing Hypomorphic Allele of Essential Protein Kinases

Chemical genetics involves the substitution of a gatekeeper residue with an amino acid containing a different side chain at the target locus. Here, we have generated a mutant parasite containing a hypomorphic allele of cdpk1 and identified compensatory pathways adopted by the parasite in the mutant background.

One of the mechanisms for subverting the effect of drugs by&#160;the malaria parasite is through rewiring of its transcriptome. The effect is more ...

Nucleofection of Parasite Forms: An Electroporation-Based Transfection Method to Deliver Fluorescent Protein Encoding-Plasmids Into Sporozoite Nucleus

Source: Duan, C., et al. <a href="https://www.jove.com/v/60552">Nucleofection and In Vivo Propagation of Chicken Eimeria Parasites.</a> J. Vis. Exp. (2020)
In this video, we describe nucleofection, an electroporation-based transfection procedure to deliver fluorescent reporter protein-encoding plasmid DNA into sporozoite form of the chicken Eimeria parasite.

Source: Duan, C., et al.  Nucleofection and In Vivo Propagation of Chicken Eimeria Parasites. J. Vis. Exp. (2020)
In this video, we describe ...

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Biological Techniques

Gene Transfer Techniques

Transfection

Intradermal Immunization with Live Attenuated Sporozoites in a Mouse Model

Source: Mac-Daniel, L., et al. <a href="https://app.jove.com/v/53796">Myeloid Cell Isolation from Mouse Skin and Draining Lymph Node Following Intradermal Immunization with Live Attenuated Plasmodium Sporozoites.</a> J. Vis. Exp. (2016).
This video demonstrates an assay for immunizing a mouse model with live attenuated sporozoites. The radiation-attenuated Plasmodium sporozoites are injected into the dorsal side of the mouse&#39;s ear. After injections at multiple locations on both ears, the mouse is allowed to recover.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

Generating Genetically Modified Plasmodium berghei Sporozoites

Summary

Explore More Videos

Chapters in this video

Protocol for Production of a Genetic Cross of the Rodent Malaria Parasites

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

Myeloid Cell Isolation from Mouse Skin and Draining Lymph Node Following Intradermal Immunization with Live Attenuated Plasmodium Sporozoites

Phenotypic Analysis of Rodent Malaria Parasite Asexual and Sexual Blood Stages and Mosquito Stages

In Vivo Tracking of Edema Development and Microvascular Pathology in a Model of Experimental Cerebral Malaria Using Magnetic Resonance Imaging

Plasmodium falciparum Gametocyte Culture and Mosquito Infection Through Artificial Membrane Feeding

Detection of Plasmodium Sporozoites in Anopheles Mosquitoes using an Enzyme-linked Immunosorbent Assay

Understanding the Development of Compensatory Pathways in a Mutant Malaria Parasite Harbouring Hypomorphic Allele of Plant-Like Kinases

Nucleofection of Parasite Forms: An Electroporation-Based Transfection Method to Deliver Fluorescent Protein Encoding-Plasmids Into Sporozoite Nucleus

Intradermal Immunization with Live Attenuated Sporozoites in a Mouse Model