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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>30 G x 1/2&#34; Syringe needle</td>
			<td>Exel international</td>
			<td>26437</td>
			<td></td>
		</tr>
		<tr>
			<td>Alsever&#39;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 dH2O 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 ...

generating-genetically-modified-plasmodium-berghei-sporozoites

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

Selection of Plasmodium falciparum Parasites for Cytoadhesion to Human Brain Endothelial Cells

Most human malaria deaths are caused by blood-stage Plasmodium falciparum parasites. Cerebral malaria, the most life-threatening complication of the disease, is characterised by an accumulation of Plasmodium falciparum infected red blood cells (iRBC) at pigmented trophozoite stage in the microvasculature of the brain2-4. This microvessel obstruction (sequestration) leads to acidosis, hypoxia and harmful inflammatory cytokines (reviewed in 5). Sequestration is also found in most microvascular tissues of the human body2, 3. The mechanism by which iRBC attach to the blood vessel walls is still poorly understood.
The immortalized Human Brain microvascular Endothelial Cell line (HBEC-5i) has been used as an in vitro model of the blood-brain barrier6. However, Plasmodium falciparum iRBC attach only poorly to HBEC-5i in vitro, unlike the dense sequestration that occurs in cerebral malaria cases. We therefore developed a panning assay to select (enrich) various P. falciparum strains for adhesion to HBEC-5i in order to obtain populations of high-binding parasites, more representative of what occurs in vivo.
A sample of a parasite culture (mixture of iRBC and uninfected RBC) at the pigmented trophozoite stage is washed and incubated on a layer of HBEC-5i grown on a Petri dish. After incubation, the dish is gently washed free from uRBC and unbound iRBC. Fresh uRBC are added to the few iRBC attached to HBEC-5i and incubated overnight. As schizont stage parasites burst, merozoites reinvade RBC and these ring stage parasites are harvested the following day. Parasites are cultured until enough material is obtained (typically 2 to 4 weeks) and a new round of selection can be performed. Depending on the P. falciparum strain, 4 to 7 rounds of selection are needed in order to get a population where most parasites bind to HBEC-5i. The binding phenotype is progressively lost after a few weeks, indicating a switch in variant surface antigen gene expression, thus regular selection on HBEC-5i is required to maintain the phenotype.
In summary, we developed a selection assay rendering P. falciparum parasites a more "cerebral malaria adhesive" phenotype. We were able to select 3 out of 4 P. falciparum strains on HBEC-5i. This assay has also successfully been used to select parasites for binding to human dermal and pulmonary endothelial cells. Importantly, this method can be used to select tissue-specific parasite populations in order to identify candidate parasite ligands for binding to brain endothelium. Moreover, this assay can be used to screen for putative anti-sequestration drugs7.

Selection of Plasmodium falciparum Parasites for Cytoadhesion to Human Brain Endothelial Cells

An in vitro model for cerebral malaria sequestration is described1. Plasmodium falciparum infected red blood cells are selected for binding to immortalized human brain microvascular endothelial cells. The selected parasites show a distinct phenotype. The selection process can be applied using various P. falciparum strains and endothelial cell lines.

Most human malaria deaths are caused by blood-stage Plasmodium falciparum parasites. Cerebral malaria, the most life-threatening complication of the ...

An In vitro Co-infection Model to Study Plasmodium falciparum-HIV-1 Interactions in Human Primary Monocyte-derived Immune Cells

Plasmodium falciparum, the causative agent of the deadliest form of malaria, and human immunodeficiency virus type-1 (HIV-1) are among the most important health problems worldwide, being responsible for a total of 4 million deaths annually1. Due to their extensive overlap in developing regions, especially Sub-Saharan Africa, co-infections with malaria and HIV-1 are common, but the interplay between the two diseases is poorly understood. Epidemiological reports have suggested that malarial infection transiently enhances HIV-1 replication and increases HIV-1 viral load in co-infected individuals2,3. Because this viremia stays high for several weeks after treatment with antimalarials, this phenomenon could have an impact on disease progression and transmission.
The cellular immunological mechanisms behind these observations have been studied only scarcely. The few in vitro studies investigating the impact of malaria on HIV-1 have demonstrated that exposure to soluble malarial antigens can increase HIV-1 infection and reactivation in immune cells. However, these studies used whole cell extracts of P. falciparum schizont stage parasites and peripheral blood mononuclear cells (PBMC), making it hard to decipher which malarial component(s) was responsible for the observed effects and what the target host cells were4,5. Recent work has demonstrated that exposure of immature monocyte-derived dendritic cells to the malarial pigment hemozoin increased their ability to transfer HIV-1 to CD4+ T cells6,7, but that it decreased HIV-1 infection of macrophages8. To shed light on this complex process, a systematic analysis of the interactions between the malaria parasite and HIV-1 in different relevant human primary cell populations is critically needed.
Several techniques for investigating the impact of HIV-1 on the phagocytosis of micro-organisms and the effect of such pathogens on HIV-1 replication have been described. We here present a method to investigate the effects of P. falciparum-infected erythrocytes on the replication of HIV-1 in human primary monocyte-derived macrophages. The impact of parasite exposure on HIV-1 transcriptional/translational events is monitored by using single cycle pseudotyped viruses in which a luciferase reporter gene has replaced the Env gene while the effect on the quantity of virus released by the infected macrophages is determined by measuring the HIV-1 capsid protein p24 by ELISA in cell supernatants.

An In vitro Co-infection Model to Study Plasmodium falciparum-HIV-1 Interactions in Human Primary Monocyte-derived Immune Cells

We have developed an in vitro malaria-HIV-1 co-infection model to study the impact of Plasmodium falciparum on the HIV-1 replicative cycle in human primary monocyte-derived macrophages. This versatile system can easily be adapted to other primary cell types susceptible to HIV-1 infection.

Plasmodium falciparum, the causative agent  of the deadliest form of malaria, and human immunodeficiency virus type-1 (HIV-1) are among the most important ...

Isolation and Analysis of Brain-sequestered Leukocytes from Plasmodium berghei ANKA-infected Mice

We describe a method for isolation and characterization of adherent inflammatory cells from brain blood vessels of P. berghei ANKA-infected mice. Infection of susceptible mouse-strains with this parasite strain results in the induction of experimental cerebral malaria, a neurologic syndrome that recapitulates certain important aspects of Plasmodium falciparum-mediated severe malaria in humans 1,2 . Mature forms of blood-stage malaria express parasitic proteins on the surface of the infected erythrocyte, which allows them to bind to vascular endothelial cells. This process induces obstructions in blood flow, resulting in hypoxia and haemorrhages 3 and also stimulates the recruitment of inflammatory leukocytes to the site of parasite sequestration.
Unlike other infections, i.e neutrotopic viruses4-6, both malaria-parasitized red blood cells (pRBC) as well as associated inflammatory leukocytes remain sequestered within blood vessels rather than infiltrating the brain parenchyma. Thus to avoid contamination of sequestered leukocytes with non-inflammatory circulating cells, extensive intracardial perfusion of infected-mice prior to organ extraction and tissue processing is required in this procedure to remove the blood compartment. After perfusion, brains are harvested and dissected in small pieces. The tissue structure is further disrupted by enzymatic treatment with Collagenase D and DNAse I. The resulting brain homogenate is then centrifuged on a Percoll gradient that allows separation of brain-sequestered leukocytes (BSL) from myelin and other tissue debris. Isolated cells are then washed, counted using a hemocytometer and stained with fluorescent antibodies for subsequent analysis by flow cytometry.
This procedure allows comprehensive phenotypic characterization of inflammatory leukocytes migrating to the brain in response to various stimuli, including stroke as well as viral or parasitic infections. The method also provides a useful tool for assessment of novel anti-inflammatory treatments in pre-clinical animal models.

Isolation and Analysis of Brain-sequestered Leukocytes from Plasmodium berghei ANKA-infected Mice

A method for isolation of adherent inflammatory leukocytes from brain blood vessels of Plasmodium berghei ANKA-infected mice is described. The method allows quantification as well as phenotypic characterization of isolated leukocytes after staining with fluorescent antibodies and subsequent analysis by flow cytometry.

We describe a method for isolation and  characterization of adherent inflammatory cells from brain blood vessels of P. berghei ANKA-infected mice. ...

Separation of Plasmodium falciparum Late Stage-infected Erythrocytes by Magnetic Means

Unlike other Plasmodium species, P. falciparum can be cultured in the lab, which facilitates its study 1. While the parasitemia achieved can reach the &asymp;40% limit, the investigator usually keeps the percentage at around 10%. In many cases it is necessary to isolate the parasite-containing red blood cells (RBCs) from the uninfected ones, to enrich the culture and proceed with a given experiment. 
When P. falciparum infects the erythrocyte, the parasite degrades and feeds from haemoglobin 2, 3. However, the parasite must deal with a very toxic iron-containing haem moiety 4, 5. The parasite eludes its toxicity by transforming the haem into an inert crystal polymer called haemozoin 6, 7. This iron-containing molecule is stored in its food vacuole and the metal in it has an oxidative state which differs from the one in haem 8. The ferric state of iron in the haemozoin confers on it a paramagnetic property absent in uninfected erythrocytes. As the invading parasite reaches maturity, the content of haemozoin also increases 9, which bestows even more paramagnetism on the latest stages of P. falciparum inside the erythrocyte.
Based on this paramagnetic property, the latest stages of P. falciparum infected-red blood cells can be separated by passing the culture through a column containing magnetic beads. These beads become magnetic when the columns containing them are placed on a magnet holder. Infected RBCs, due to their paramagnetism, will then be trapped inside the column, while the flow-through will contain, for the most part, uninfected erythrocytes and those containing early stages of the parasite.
Here, we describe the methodology to enrich the population of late stage parasites with magnetic columns, which maintains good parasite viability 10. After performing this procedure, the unattached culture can be returned to an incubator to allow the remaining parasites to continue growing.

Separation of Plasmodium falciparum Late Stage-infected Erythrocytes by Magnetic Means

The paramagnetic properties of hemozoin are used to isolate late stages of Plasmodium falciparum-infected red blood cells growing in culture. The method is simple and fast and does not affect the subsequent invasive capabilities of the parasites.

Unlike other Plasmodium species, P. falciparum can be cultured in the lab, which facilitates its study 1. While the parasitemia achieved can reach the ...

An Experimental Model to Study Tuberculosis-Malaria Coinfection upon Natural Transmission of Mycobacterium tuberculosis and Plasmodium berghei

Coinfections naturally occur due to the geographic overlap of distinct types of pathogenic organisms. Concurrent infections most likely modulate the respective immune response to each single pathogen and may thereby affect pathogenesis and disease outcome. Coinfected patients may also respond differentially to anti-infective interventions. Coinfection between tuberculosis as caused by mycobacteria and the malaria parasite Plasmodium, both of which are coendemic in many parts of sub-Saharan Africa, has not been studied in detail. In order to approach the challenging but scientifically and clinically highly relevant question how malaria-tuberculosis coinfection modulate host immunity and the course of each disease, we established an experimental mouse model that allows us to dissect the elicited immune responses to both pathogens in the coinfected host. Of note, in order to most precisely mimic naturally acquired human infections, we perform experimental infections of mice with both pathogens by their natural routes of infection, i.e. aerosol and mosquito bite, respectively.

An Experimental Model to Study Tuberculosis-Malaria Coinfection upon Natural Transmission of Mycobacterium tuberculosis and Plasmodium berghei

Tuberculosis and malaria are two of the most prevalent infections in humans and major causes of morbidity and mortality in impoverished populations in the tropics. We established an experimental model system to study outcome of malaria-tuberculosis coinfection in mice after challenge with both pathogens via their natural route of infection.

Coinfections naturally occur due to the geographic overlap of distinct types of pathogenic organisms. Concurrent infections most likely modulate the ...

A Method for Generating Pulmonary Neutrophilia Using Aerosolized Lipopolysaccharide

Acute lung injury (ALI) is a severe disease characterized by alveolar neutrophilia, with limited treatment options and high mortality. Experimental models of ALI are key in enhancing our understanding of disease pathogenesis. Lipopolysaccharide (LPS) derived from gram positive bacteria induces neutrophilic inflammation in the airways and lung parenchyma of mice. Efficient pulmonary delivery of compounds such as LPS is, however, difficult to achieve. In the approach described here, pulmonary delivery in mice is achieved by challenge to aerosolized Pseudomonas aeruginosa LPS. Dissolved LPS was aerosolized by a nebulizer connected to compressed air. Mice were exposed to a continuous flow of LPS aerosol in a Plexiglas box for 10 min, followed by 2 min conditioning after the aerosol was discontinued. Tracheal intubation and subsequent bronchoalveolar lavage, followed by formalin perfusion was next performed, which allows for characterization of the sterile pulmonary inflammation. Aerosolized LPS generates a pulmonary inflammation characterized by alveolar neutrophilia, detected in bronchoalveolar lavage and by histological assessment. This technique can be set up at a small cost with few appliances, and requires minimal training and expertise. The exposure system can thus be routinely performed at any laboratory, with the potential to enhance our understanding of lung pathology.

We describe a method for inducing neutrophilic pulmonary inflammation by challenge to aerosolized lipopolysaccharide by nebulization, to model acute lung injury. In addition, basic surgical techniques for lung isolation, tracheal intubation and bronchoalveolar lavage are also described.

Acute lung injury (ALI) is a severe disease characterized by alveolar neutrophilia, with limited treatment options and high mortality. Experimental models ...

High Yield Purification of Plasmodium falciparum Merozoites For Use in Opsonizing Antibody Assays

Plasmodium falciparum merozoite antigens are under development as potential malaria vaccines. One aspect of immunity against malaria is the removal of free merozoites from the blood by phagocytic cells. However assessing the functional efficacy of merozoite specific opsonizing antibodies is challenging due to the short half-life of merozoites and the variability of primary phagocytic cells. Described in detail herein is a method for generating viable merozoites using the E64 protease inhibitor, and an assay of merozoite opsonin-dependent phagocytosis using the pro-monocytic cell line THP-1. E64 prevents schizont rupture while allowing the development of merozoites which are released by filtration of treated schizonts. &#160;Ethidium bromide labelled merozoites are opsonized with human plasma samples and added to THP-1 cells. Phagocytosis is assessed by a standardized high throughput protocol. Viable merozoites are a valuable resource for assessing numerous aspects of P. falciparum biology, including assessment of immune function. Antibody levels measured by this assay are associated with clinical immunity to malaria in naturally exposed individuals. The assay may also be of use for assessing vaccine induced antibodies. &#160;

High Yield Purification of Plasmodium falciparum Merozoites For Use in Opsonizing Antibody Assays

Measuring antibody function is key to understanding immunity to Plasmodium falciparum malaria. This method describes the purification of viable merozoites, and measurement of opsonization-dependent phagocytosis by flow cytometry.

Plasmodium falciparum merozoite antigens are under development as potential malaria vaccines. One aspect of immunity against malaria is the removal of ...

In Vivo Assessment of Rodent Plasmodium Parasitemia and Merozoite Invasion by Flow Cytometry

During blood stage infection, malaria parasites invade, mature, and replicate within red blood cells (RBCs). This results in a regular growth cycle and an exponential increase in the proportion of malaria infected RBCs, known as parasitemia. We describe a flow cytometry based protocol which utilizes a combination of the DNA dye Hoechst, and the mitochondrial membrane potential dye, JC-1, to identify RBCs which contain parasites and therefore the parasitemia, of in vivo blood samples from Plasmodium chabaudi adami DS infected mice. Using this approach, in combination with fluorescently conjugated antibodies, parasitized RBCs can be distinguished from leukocytes, RBC progenitors, and RBCs containing Howell-Jolly bodies (HJ-RBCs), with a limit of detection of 0.007% parasitemia. Additionally, we outline a method for the comparative assessment of merozoite invasion into two different RBC populations. In this assay RBCs, labeled with two distinct compounds identifiable by flow cytometry, are transfused into infected mice. The relative rate of invasion into the two populations can then be assessed by flow cytometry based on the proportion of parasitized RBCs in each population over time. This combined approach allows the accurate measurement of both parasitemia and merozoite invasion in an in vivo model of malaria infection.

In Vivo Assessment of Rodent Plasmodium Parasitemia and Merozoite Invasion by Flow Cytometry

The malaria parasite invades and replicates within red blood cells. The accurate assessment of merozoite invasion and parasitemia is therefore crucial in assessing the course of malaria infection. Here we describe a flow cytometry based protocol for the measurement of these parameters in a mouse model of malaria.

During blood stage infection, malaria parasites invade, mature, and replicate within red blood cells (RBCs). This results in a regular growth cycle and an ...

Methods to Investigate the Regulatory Role of Small RNAs and Ribosomal Occupancy of Plasmodium falciparum

The genetic variation responsible for the sickle cell allele (HbS) enables erythrocytes to resist infection by the malaria parasite, P. falciparum. The molecular basis of this resistance, which is known to be multifactorial, remains incompletely understood. Recent studies found that the differential expression of erythrocyte microRNAs, once translocated into malaria parasites, affect both gene regulation and parasite growth. These miRNAs were later shown to inhibit mRNA translation by forming a chimeric RNA transcript via 5&#39; RNA fusion with discreet subsets of parasite mRNAs. Here, the techniques that were used to study the functional role and putative mechanism underlying erythrocyte microRNAs on the gene regulation and translational potential of P. falciparum, including the transfection of modified synthetic microRNAs into host erythrocytes, will be detailed.&#160; Finally, a polysome gradient method is used to determine the extent of translation of these transcripts. Together, these techniques allowed us to demonstrate that the dysregulated levels of erythrocyte microRNAs contribute to cell-intrinsic malaria resistance of sickle erythrocytes.

Methods to Investigate the Regulatory Role of Small RNAs and Ribosomal Occupancy of Plasmodium falciparum

Human microRNAs translocate from host erythrocytes to Plasmodium falciparum parasites. Here, the techniques used to transfect synthetic microRNAs into host erythrocytes and isolate all RNAs from P. falciparum are described. In addition, this paper will detail a method of polysome isolation in P. falciparum to determine the ribosomal occupancy and translational potential of parasite transcripts.

The genetic variation responsible for the sickle cell allele (HbS) enables erythrocytes to resist infection by the malaria parasite, P. falciparum. The ...

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 ...