We thank Timothy J. Kurtti and Benjamin Cull for their insightful discussions and suggestions. This study was financially supported by a grant to U.G.M. from the NIH (2R01AI049424) and a grant to U.G.M. from the Minnesota Agricultural Experiment Station (MIN-17-078).

1. Propagation and purification of R. parkeri from tick cell culture
NOTE: All cell culture procedures are to be performed in a class II biosafety cabinet.
<ol>
	<li>Preparing R. parkeri-infected tick cells
	<ol>
		<li>Grow ISE6 cells in 25 cm2 cell culture flasks at 34 &#176;C in 5 mL of L15C300 medium17, supplemented with 5% fetal bovine serum (FBS), 5% tryptose phosphate broth (TPB), and 0.1% lipoprotein concentrate (LPC). 
		NOTE: ISE6 (Ixodes scapularis embryo-derived cell line) is a tick cell line extensively used in many laboratories and is, thus, an essential model to study tick-pathogen interactions17,18,19. The growth rate of ISE6 cells is slower than that of mammalian cells, with a population doubling time of &#8805;72 h. For example, ISE6 cells subcultivated from a 100% confluent culture at a ratio of 1:5 will take 3-4 weeks to regain 100% confluency. Healthy ISE6 cells will be generally rounded with numerous adhering filaments17.</li>
		<li>Count the ISE6 cells with a hemocytometer under a light microscope. Use at a concentration of ~106 cells/mL (100% confluent).
		<ol>
			<li>Gently rinse the cells from the flask with medium and disperse the cells by pipetting to generate a homogenous cell suspension. Add 20 &#181;L of cell suspension between the hemocytometer and cover glass to count the cells using a hemocytometer.</li>
		</ol>
		</li>
		<li>Add host cell-free wild-type R. parkeri20&#160;(average number: 5 &#215; 107-10 &#215; 107) to the ISE6 cell cultures in 25 cm2 cell culture flasks. Incubate infected R. parkeri cell cultures at 34 &#176;C in 5 mL of complete medium consisting of L15C300 medium, 10% FBS, 5% TPB, 0.1% LPC, 0.25% sodium bicarbonate (NaHCO3), and 25 mM HEPES until 90%-100% of the cells are infected. 
		NOTE: The infection rate of R. parkeri in ISE6 cells is determined by Giemsa staining, and the incubation time to reach 90%-100% infection ranges from 5 days to 7 days on average.</li>
		<li>Determine the infection rate via Giemsa staining.
		<ol>
			<li>In the biosafety cabinet, resuspend the infected cell cultures and transfer 50 &#181;L of the suspension to a 1.5 mL microcentrifuge tube.</li>
			<li>Dilute the suspension with complete medium (1:5 dilution) and centrifuge 100 &#181;L onto glass microscope slides using a centrifuge at 113 &#215; g for 5 min. To keep live R. parkeri contained, use a centrifuge with a sealed rotor that is designed to be removed from the centrifuge. This enables the transport of the sealed rotor in and out of the hood. 
			NOTE: As the R. parkeri are still alive before centrifugation, the sample needs to be contained until the rickettsia are killed. Thus, the centrifuge used to spin the R. parkeri culture onto the slide must have a sealed rotor that is designed to be lifted out of the centrifuge. With the rotor in the laminar flow hood, load the samples into the funnel-microscope slide assembly, reseal the rotor, and place the sealed rotor back into the centrifuge. Then, turn on the centrifuge, and the samples are deposited onto the microscope slides. After the run is finished, remove the sealed rotor from the centrifuge and place it in the laminar flow hood. There, open the rotor lid, and unload the funnel-microscope slide assembly inside the hood. Once dry, immerse the glass microscope slides in absolute methanol, which kills all pathogens. The funnels and the metal carriers are submerged in diluted DMQ, which kills R. parkeri.</li>
			<li>Air-dry the slides and fix in absolute methanol for 5 min at room temperature (RT).</li>
			<li>Stain the slides with Giemsa stain (4% in S&#248;renson&#39;s buffer, pH 6.6) for 30 min at 37 &#176;C and rinse with water for 5 s.</li>
			<li>Determine the infection percentage (number of cells infected with rickettsiae/100 cells) by light microscopy. 
			NOTE: Figure 1 shows typical Giemsa stain results.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Preparing cell-free R. parkeri 
	NOTE: When 90%-100% of the cells in the culture are infected, the rickettsiae should be isolated from the ISE6 cells and electroporation should be performed.
	<ol>
		<li>Add sterile 60-90 silicon carbide grit to sterile 2 mL microcentrifuge tubes to a volume of approximately 0.2 mL.</li>
		<li>Remove 2 mL of medium from 100% R. parkeri-infected cultures (step 1.1.3) and resuspend the cells in the remaining 3 mL of medium.</li>
		<li>Transfer equal portions of this suspension to tubes prepared in step 1.2.1.</li>
		<li>Vortex each tube at high speed for 30 s and then place the tubes on ice. Let the grit settle within seconds by gravity.</li>
		<li>In a class II biosafety cabinet, have a sterile 5 mL Luer-lock syringe ready with the plunger extended and the end of the plunger secured in a polystyrene base for 15 mL conical tubes.</li>
		<li>Using a sterile, 1 mL barrier pipet tip fitted to a suitable pipettor, remove the supernatant of the vortexed cell lysate from the tube, taking care not to aspirate any grit. Insert the pipet tip into the syringe hub opening and eject the contents into the syringe with gentle pressure. Alternatively, use a sterile, barrier 2 mL Pasteur pipette operated with a rubber bulb.</li>
		<li>Attach a sterilized 2 &#181;m pore size filter onto the syringe hub and filter the R. parkeri culture from step 1.2.6 into sterile 1.5 mL tubes.</li>
		<li>Collect the R. parkeri by centrifugation at 13,600 &#215; g at 4 &#176;C for 5 min, and discard the supernatant.</li>
		<li>Resuspend the pellet in 1.2 mL of ice-cold 300 mM sucrose and centrifuge again at 13,600 &#215; g at 4 &#176;C for 5 min. Repeat the resuspension and centrifugation for a total of two sucrose washes.</li>
		<li>Combine the R. parkeri pellets into one chilled, sterile 1.5 mL microcentrifuge tube in 50 &#181;L of ice-cold 300 mM sucrose per transformation. As one 25 cm2 flask of R. parkeri provides enough rickettsiae for two to three transformations, add 100-150 &#181;L of 300 mM cold sucrose. 
		NOTE: If desired, the number of rickettsiae can be calculated using a Petroff-Hausser counting chamber. When using an initial inoculation of 5 &#215; 107 -10 &#215; 107 cell-free R. parkeri, it will take 5-7 days on average to reach 90%-100% infection, yielding approximately 5 &#215; 107-10 &#215; 1010 cell-free R. parkeri.</li>
		<li>Gently resuspend the pellet by pipetting until it is thoroughly dispersed. Divide the sample into 50 &#181;L portions in chilled, sterile 1.5 mL microcentrifuge tubes. 
		NOTE: If desired, rickettsial viability may be assessed by flow cytometry following staining with a fluorescent dye 21.</li>
	</ol>
	</li>
</ol>
2. Transformation of R. parkeri with the pRAM18dSFA plasmid
<ol>
	<li>On ice, add 3 &#181;g of endotoxin-free pRAM18dSFA plasmid DNA13,15,20 to each tube containing 50 &#181;L of R. parkeri suspension and stir the mixture with the pipet tip gently but thoroughly. 
	NOTE: When preparing plasmid DNA, use a purification kit that includes an endotoxin-removal step to yield endotoxin-free plasmid DNA23. We found that a range of plasmid concentrations of between 1 &#181;g and 3 &#181;g per 50 &#181;L of R. parkeri suspension resulted in successful transformations, whereas increasing the amount of plasmid to 10 &#181;g inhibited transformation.</li>
	<li>Transfer the above mixture of DNA and R. parkeri into a chilled, sterile 0.1 cm gap electroporation cuvette (gently tap the cuvette until the mixture is evenly distributed) and let it sit on ice for 10-30 min.</li>
	<li>Remove the medium from a 100% confluent culture of ISE6 cells and resuspend the cell layers in 1.5 mL of fresh medium with NaHCO3 and HEPES buffer (described above in step 1.1.3). Use one flask of confluent cells per transformation.</li>
	<li>Transfer the resuspended cells to a sterile 2 mL microcentrifuge tube (one tube per 25 cm2 flask of ISE6 cells).</li>
	<li>Electroporate the R. parkeri/pRAM18dSFA mixture at 1.8 KV, 200 ohms, 25 &#181;F using an electroporator.</li>
	<li>Using a sterile, extended fine-tip transfer pipet, transfer a small amount of the ISE6 cell suspension (step 2.4) into the cuvette and gently pull the liquid up and down to wash out the cuvette. Transfer the mixture into the 2 mL microcentrifuge tube containing the remainder of the ISE6 cell suspension and gently mix the transformation mixture with the cells.</li>
	<li>Centrifuge the transformed samples at 700 &#215; g for 2 min at RT, and then increase the centrifugation speed to 13,600 &#215; g for 1 min more at RT. 
	NOTE: The two centrifugation speeds promote rickettsial binding with and entry into the cells: 700 &#215; g is used to pull down the ISE6 cells, while 13,600 &#215; g is used to pull down the rickettsiae.</li>
	<li>Let samples sit at RT or 34 &#176;C for 15 min to 1 h.</li>
	<li>Resuspend the transformants (two or three) in ISE6 cells with a sterile 2 mL barrier pipet tip fitted to a pipettor and transfer into two or three 25 cm2 cell culture flasks containing 3.5 mL of fresh medium with NaHCO3 and HEPES buffer. Alternatively, use a sterile, barrier 2 mL Pasteur pipette operated with a rubber bulb.</li>
	<li>Rock the flasks to spread the mixture evenly and then incubate at 34 &#176;C.</li>
	<li>After 16-24 h, add 10 &#181;L of spectinomycin (50 mg/mL) and 10 &#181;L of streptomycin (50 mg/mL) to each flask in step 2.10.</li>
</ol>
3. Observation of the transformed R. parkeri
NOTE: Use an epifluorescence microscope with rhodamine/TRITC filters to observe the flasks prepared in step 2.11 after 3-7 days. Once plaques are evident in the cultures (5-14 days), transformed R. parkeri can be seen that express the red fluorescent protein mKATE, encoded on the pRAM18dSFA plasmid.
<ol>
	<li>Confocal microscopy
	<ol>
		<li>Resuspend the cell cultures from the step 2.11 flasks and mix 100 &#181;L of these cell cultures with 5 &#181;L of Hoechst 33342 solution for 10-30 min at RT in the dark. 
		NOTE: Hoechst 33342 is a cell-permeable nuclear counterstain that binds to live tick cell DNA and emits blue fluorescence.</li>
		<li>Centrifuge 50 &#181;L of the mixture onto glass microscope slides using a centrifuge at 5 &#215; g for 3 min.</li>
		<li>Add 3 &#181;L of 1x PBS to the spot of cells deposited on the slide, overlay with a cover slip, and view with a confocal microscope (60x objective). Use the following excitation and emission parameters for fluorescent imaging: for 4&#8242;,6-diamidino-2-phenylindole (DAPI), excitation at 350 nm, emission at 470 nm; for tetramethylrhodamine (TRITC), excitation at 557 nm, emission at 576 nm.</li>
	</ol>
	</li>
</ol>

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No conflicts of interest are declared.

Here, we demonstrate a method for introducing exogenous DNA encoded on the shuttle plasmid pRAM18dSFA into rickettsiae using electroporation. In this procedure, cell-free rickettsiae were purified from host cells, transformed with a rickettsial shuttle vector, and released onto tick cells for infection. Also described is a confocal immunofluorescence procedure to detect red fluorescence protein-expressing R. parkeri in tick cells. Similar methods are applicable to other Rickettsia species and with further modifications might be adaptable to other obligate intracellular bacteria that are capable of maintaining a plasmid.
A successful transformation in this protocol requires three components: exogenous DNA, Rickettsia spp. bacteria, and a suitable type of host cell22. Depending on the specific research goal, these components can be changed. First, the exogenous DNA was introduced via the pRAM18dSFA plasmid, which contains spectinomycin- and streptomycin-selectable markers and expresses mKATE (a far-red fluorescent protein). This plasmid also confers ampicillin resistance for growth in Escherichia coli. As shown in previous studies, it is possible to replace antibiotic-resistance markers and the genes for fluorescent proteins13. Second, ISE6 cells were selected to represent transformed cell lines as this cell line is extensively used in many laboratories and is an essential model for the study of vector-pathogen interactions17,18,19. Other species of tick16 or mammalian cells23,24 can also be used to grow rickettsiae for transformation. Finally, in this protocol, R. parkeri was used, which can be manipulated in a biosafety level 2 (BSL-2) laboratory using a class II biosafety cabinet with proper safety precautions. Other more pathogenic Rickettsia spp. (e.g., R. rickettsii; R. prowazekii) require a BSL-3 setting for manipulation; therefore, a more stringent biosafety standard protocol must be used when working with such agents.
Although this protocol can be used as a standard procedure for rickettsiae transformation, several key technical steps require particular attention. First, a successful purification step must ensure cell-free rickettsiae survival and infectivity9; hence, the most critical aspect of the transformation procedure is to isolate infectious, cell-free rickettsiae. Any residual salt in the purified rickettsiae will result in arcing and transformation failure. Therefore, washing the isolated cell-free rickettsiae with sucrose is required to remove all salts. The cold sucrose solution also protects the rickettsial membrane integrity in the extracellular condition.
When 90%-100% of the cells in a culture are infected, the rickettsiae should be isolated from the ISE6 cells, and electroporation should be performed without delay and without interruptions, as rickettsiae are intracellular organisms and do not survive well outside cells. Although rickettsial cultures can be stored at 4 &#176;C, this type of material should not be used for electroporation. A culture that has been stored at 4 &#176;C can be used to inoculate a fresh layer of ISE6 cells, which can then be used for electroporation once infection has reached 90%-100%.
Second, the electroporation settings, including voltage, resistance, capacitance, and time constant, are specific to different rickettsial species. For example, the field strength required during electroporation is dependent on the size of the rickettsiae and exogenous DNA7. Finally, it is important to maintain the transformed rickettsiae under antibiotic selection to prevent the loss of exogenous plasmids during multiple passages in host cells9. The concentration and type of antibiotic employed are dependent on the Rickettsia species being transformed, as previously described13,15,16,23, and it is important that the antibiotic marker selected should not be one that is applied in clinical treatments.
To follow this protocol, both spectinomycin and streptomycin should be used for selection until all the residual untransformed rickettsiae have been eliminated. Combined, the two antibiotics kill both intracellular and extracellular rickettsiae and reduce the probability of resistant wild-type rickettsiae emerging. In addition, using both of these antibiotics does not affect downstream experiments, such as the ability to use them separately to select for two different plasmids, since the aadA gene confers resistance to both spectinomycin and streptomycin. The two antibiotics used in this protocol do not confer resistance to antibiotics used in the clinical treatment of rickettsial disease.
The ISE6 tick cell line utilized in this study has unique advantages. First, ISE6 cells were isolated from the black-legged tick Ixodes scapularis Say (Acari: Ixodidae), the principal vector for seven human pathogens in the United States. Second, ISE6 is an extensively used tick cell line in many laboratories and has been successfully used to recover many tick-associated bacteria (Rickettsia, Anaplasma, and Ehrlichia) following electroporation. Third, some pathogenic rickettsiae can only be propagated in tick cells but not in mammalian cells17,18,19,24. However, tick cell lines are relatively fragile compared to mammalian cells and have more intensive culture requirements25,26,27,28. In addition, the growth rate of ISE6 cells is significantly slower than that of mammalian cell lines, even if seeded at the same initial density, although slow replication can be advantageous when the rickettsial transformants exhibit reduced growth.
This protocol also provides a method to evaluate the transformation efficiency of rickettsiae in live cells at critical experimental steps, which helps to optimize electroporation settings or in testing the efficiency of various cell lines for recovering transformants. Nevertheless, this protocol has limitations, especially for the quantification of the transformants obtained. Transformation efficiency as an indicator of a successful transformation could be demonstrated in a future study. Rickettsial viability could be assessed using a suitable staining kit and flow cytometry21 to quantify the transformants that are obtained. Two parameters would be used to determine transformation efficiency-the number of live rickettsiae used for transformation and the number of transformed rickettsiae that express mKATE.
In addition, image analysis with fluorescence signals could be used to compare the transformation rates of different plasmids. As the underlying mechanism of rickettsial plasmid maintenance is poorly understood, well-characterized transformation systems for introducing exogenous plasmids can be valuable tools for further investigation. Furthermore, the direct visualization of fluorescent protein-expressing rickettsiae in cells or tissues will improve our understanding of rickettsia/host/vector interactions. This will provide information for designing strategies to control and prevent rickettsioses.
Data availability: 
All data underlying the results of this study are publicly available.

Rickettsioses are caused by a broad range of obligate intracellular bacteria that belong to the genus Rickettsia (family Rickettsiaceae, order Rickettsiales). The genus Rickettsia is classified into four major groups based on phylogenetic characteristics1,2: the spotted fever group (SFG), which contains those rickettsiae that cause the most severe and fatal tick-borne rickettsioses (e.g., Rickettsia rickettsii, the causative agent of Rocky Mountain Spotted Fever), the typhus group (TG, e.g., Rickettsia prowazekii, the agent of epidemic typhus), the transitional group (TRG, e.g., Rickettsia felis, the causative agent of flea-borne spotted fever), and the ancestral group (AG, e.g., Rickettsia bellii).
Among the oldest known vector-borne diseases, rickettsioses are mainly acquired following transmission of the pathogens through the bites of infected arthropod vectors, including ticks, fleas, lice, and mites3,4. Although the discovery of effective antibiotics improved treatment outcomes, emerging and re-emerging epidemic rickettsioses continue to challenge traditional prevention and control strategies. Thus, a comprehensive understanding of rickettsia/host/vector interactions would ultimately establish a strong foundation for developing new approaches to prevent and cure these ancient diseases.
In nature, horizontal gene transfer (HGT) in bacteria occurs through conjugation, transduction, and transformation5. In vitro bacterial transformation utilizes these HGT concepts, although the intracellular nature of rickettsiae presents some challenges. The restricted growth conditions and poorly understood conjugation and transduction systems in different species of rickettsiae have prevented the application of conjugation and transduction methods in rickettsiae6,7,8. Compared with other obligate intracellular bacterial genera (e.g., Chlamydia, Coxiella, Anaplasma, and Ehrlichia), the genus Rickettsia differs with regard to the growth and replication strategies within the cell cytoplasm, which imposes specific challenges to the genetic modification of rickettsiae due to their unique lifestyle features9.
The initial hurdle to overcome when attempting the genetic modification of rickettsiae is to achieve successful transformation. Thus, designing a feasible approach with high transformation efficiency would be extremely valuable for developing genetic tools for rickettsiae. Here, we focus on electroporation, a broadly recognized transformation method that has been used to introduce exogenous DNA successfully into several species of rickettsiae, including Rickettsia prowazekii, Rickettsia typhi, Rickettsia conorii, Rickettsia parkeri, Rickettsia montanensis, Rickettsia bellii, Rickettsia peacockii, and Rickettsia buchneri10,11,12,13,14,15,16.
This paper describes a procedure for the electroporation of the R. parkeri strain Tate&#39;s Hell (accession: GCA_000965145.1) with the shuttle vector pRAM18dSFA derived from the Rickettsia amblyommatis strain AaR/SC plasmid pRAM18 engineered to encode mKATE, a far-red fluorescent protein, and aadA, conferring spectinomycin and streptomycin resistance13,15,20. Transformed R. parkeri are viable and stably maintained under antibiotic selection in tick cell lines. In addition, we show that the localization of transformed R. parkeri in live tick cells via confocal microscopy can be used to assess the quality of transformation rates in vector cell lines.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.1 cm gap gene pulser electroporation cuvette</td>
			<td>Bio-Rad</td>
			<td>1652083</td>
			<td></td>
		</tr>
		<tr>
			<td>2 &#956;m pore size filter&#160;</td>
			<td>GE Healthcare Life Sciences Whatman</td>
			<td>6783-2520</td>
			<td></td>
		</tr>
		<tr>
			<td>5 mL Luer-lock syringe&#160;</td>
			<td>BD</td>
			<td>309646</td>
			<td></td>
		</tr>
		<tr>
			<td>60-90 silicon carbide grit&#160;</td>
			<td>LORTONE, inc</td>
			<td>&#160;591-056</td>
			<td></td>
		</tr>
		<tr>
			<td>absolute methanol&#160;</td>
			<td>Fisher Scientific</td>
			<td>A457-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Bacto tryptose phosphate broth&#160;</td>
			<td>BD</td>
			<td>260300</td>
			<td></td>
		</tr>
		<tr>
			<td>Cytospin centrifuge Cytospin4</td>
			<td>Thermo Fisher Scientific</td>
			<td>A78300003</td>
			<td>The rotor is detatchable so the whole rotor can be put into the hood to load infectious samples</td>
		</tr>
		<tr>
			<td>EndoFree Plasmid Maxi Kit (10)</td>
			<td>QIAGEN</td>
			<td>12362</td>
			<td>used to obtain endotoxin-free pRAM18dSFA plasmid</td>
		</tr>
		<tr>
			<td>extended fine tip transfer pipet&#160;</td>
			<td>Perfector Scientific</td>
			<td>TP03-5301</td>
			<td></td>
		</tr>
		<tr>
			<td>fetal bovine serum&#160;</td>
			<td>Gemini Bio</td>
			<td>900-108</td>
			<td>The FBS batch has to be tested to make sure ISE6 cells will grow well in it.</td>
		</tr>
		<tr>
			<td>Gene Pulser II electroporator with Pulse Controller PLUS</td>
			<td>Bio-Rad</td>
			<td>165-2105 &#38; 165-2110</td>
			<td></td>
		</tr>
		<tr>
			<td>hemocytometer</td>
			<td>Thermo Fisher Scientific</td>
			<td>267110</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Millipore-Sigma</td>
			<td>H4034</td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ Fiji</td>
			<td>National Institute of Health</td>
			<td></td>
			<td>raw image editing</td>
		</tr>
		<tr>
			<td>KaryoMAX Giemsa stain&#160;</td>
			<td>Gibco</td>
			<td>&#160;2021-10-30</td>
			<td></td>
		</tr>
		<tr>
			<td>Leibovitz&#39;s L-15 medium</td>
			<td>Gibco</td>
			<td>41300039</td>
			<td></td>
		</tr>
		<tr>
			<td>lipoprotein concentrate&#160;</td>
			<td>MP Biomedicals</td>
			<td>191476</td>
			<td></td>
		</tr>
		<tr>
			<td>Nikon Diaphot</td>
			<td>Nikon</td>
			<td></td>
			<td>epifluorescence microscope</td>
		</tr>
		<tr>
			<td>NucBlue Live ReadyProbes Reagent&#160;</td>
			<td>Thermo Fisher Scientific</td>
			<td>R37605</td>
			<td></td>
		</tr>
		<tr>
			<td>Olympus Disc Scanning Unit (DSU) confocal microscope&#160;</td>
			<td>Olympus</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Petroff-Hausser Counting Chamber</td>
			<td>Hausser Scientific</td>
			<td>&#160;Chamber 3900</td>
			<td></td>
		</tr>
		<tr>
			<td>sodium bicarbonate</td>
			<td>Millipore-Sigma</td>
			<td>S5761</td>
			<td></td>
		</tr>
		<tr>
			<td>Vortex</td>
			<td>Fisher Vortex Genie 2</td>
			<td>12-812</td>
			<td></td>
		</tr>
	</tbody>
</table>

an electroporation method to transform rickettsia spp. with a fluorescent protein-expressing shuttle vector in tick cell lines

Rickettsioses are caused by a broad range of obligate intracellular bacteria belonging to the genus Rickettsia that can be transmitted to vertebrate hosts through the bite of infected arthropod vectors. To date, emerging or re-emerging epidemic rickettsioses remain a public health risk due to the difficulty in diagnosis, as diagnostic methods are limited and not standardized or universally accessible. Misdiagnosis resulting from a lack of recognition of the signs and symptoms may result in delayed antibiotic treatment and poor health outcomes. A comprehensive understanding of Rickettsia characteristics would ultimately improve clinical diagnosis, assessment, and treatment with improved control and prevention of the disease. 
Functional studies of rickettsial genes are crucial for understanding their role in pathogenesis. This paper describes a procedure for the electroporation of the Rickettsia parkeri strain Tate's Hell with the shuttle vector pRAM18dSFA and the selection of transformed R. parkeri in tick cell culture with antibiotics (spectinomycin and streptomycin). A method is also described for the localization of transformed R. parkeri in tick cells using confocal immunofluorescence microscopy, a useful technique for checking transformation in vector cell lines. Similar approaches are also suitable for the transformation of other rickettsiae.

The morphology of R. parkeri in ISE6 cells under a light microscope after Giemsa staining are shown in Figure 1. In Figure 2, transformed R. parkeri expressing red fluorescence protein in ISE6 cells are shown using confocal microscopy. There is a substantial increase in the infection rate of transformed R. parkeri (red) in ISE6 cells (blue, corresponds to the nuclei) from (A) day 7 to (B) day 10 of incubation.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64562/64562fig01.jpg" /> 
Figure 1: Slides showing Giemsa-stained Rickettsia parkeri in ISE6 cells. ISE6 cells infected with wild-type R. parkeri at (A) low infection and (B) 90%-100% infection rates. The nuclei of the ISE6 cells stain to dark purple with Giemsa; the R. parkeri appear as dark purple rods. The red boxes show intracellular rickettsiae, and the red asterisks indicate extracellular rickettsiae. All images were taken on a light microscope with a 100x objective. Scale bars = 50 &#181;m. Abbreviation: N = nuclei. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64562/64562fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64562/64562fig02.jpg" /> 
Figure 2: Transformed Rickettsia parkeri expressing red fluorescence protein in ISE6 cells. pRAM18dSFA-transformed R. parkeri (red) in ISE6 cells (blue), stained with Hoechst 33342 and detected by confocal microscopy (A) 7 days and (B) 10 days after transformation. Hoechst 33342 stains the nuclei, and its excitation and emission wavelengths are similar to DAPI. The merged signals are a combination of DAPI and TRITC filter images. All images were taken on a confocal microscope with a 60x objective. Scale bars = 20 &#181;m. Abbreviations: DAPI = 4&#39;,6-diamidino-2-phenylindole; TRITC = tetramethylrhodamine. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64562/64562fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about An Electroporation Method to Transform Rickettsia spp. with a Fluorescent Protein-Expressing Shuttle Vector in Tick Cell Lines at JoVE.com

An Electroporation Method to Transform Rickettsia spp. with a Fluorescent Protein-Expressing Shuttle Vector in Tick Cell Lines

Electroporation is a rapid, broadly adopted method for introducing exogenous DNA into the genus Rickettsia. This protocol provides a useful electroporation method for the transformation of obligate intracellular bacteria in the genus Rickettsia.

An Electroporation Method to Transform Rickettsia spp. with a Fluorescent Protein-Expressing Shuttle Vector in Tick Cell Lines

Rickettsioses are caused by a broad range of obligate intracellular bacteria belonging to the genus Rickettsia that can be transmitted to vertebrate hosts ...

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1.7K Views.  University of Minnesota. Electroporation is a rapid, broadly adopted method for introducing exogenous DNA into the genus Rickettsia. This protocol provides a useful electroporation method for the transformation of obligate intracellular bacteria in the genus Rickettsia. 

Video: An Electroporation Method to Transform Rickettsia spp. with a Fluorescent Protein-Expressing Shuttle Vector in Tick Cell Lines

Passaging HuES Human Embryonic Stem Cell-lines with Trypsin.

In this video we demonstrate how our lab routinely passages HuES human embryonic stem cell lines with trypsin.  Human embryonic stem cells are artifacts of cell culture, and tend to acquire karyotypic abnormalities with high population doublings.  Proper passaging is essential for maintaining a healthy, undifferentiated, karyotypically normal HuES human embryonic stem cell culture.  First, an expanding culture is washed in PBS to remove residual media and cell debris, then cells are overlaid with a minimal volume of warm 0.05% Trypsin-EDTA.  Trypsin is left on the cells for up to five minutes, then cells are gently dislodged with a 2mL serological pipette.  The cell suspension is collected and mixed with a large volume of HuES media, then cells are collected by gentle centrifugation.  The inactivated trypsin media mixture is removed, and cells resuspended in pre-warmed HuES media.  An appropriate split ratio is calculated (generally 1:10 to 1:20), and cells re-plated onto a 1-2 day old plate containing a monolayer of irradiated mouse embryonic fibroblast feeder cells.  The newly seeded HuES culture plate is left undisturbed for 48 hrs, then media is changed every day thereafter.  It is important not to trpsinize down to a single cell suspension, as this increases the risk of introducing karyotypic abnormalities.

In this video we demonstrate how our lab routinely passages HuES human embryonic stem cell lines with trypsin. Brought to you by JoVE.

Barwy Pasażowanie ludzkich embrionalnych komórek macierzystych linii Trypsyna

In this video we demonstrate how our lab routinely passages HuES human embryonic stem cell lines with trypsin.  Human embryonic stem cells are artifacts ...

Electroporation of Mycobacteria

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly caused by the thick and waxy cell wall, but is compounded by the fact that most molecular techniques have been developed for distantly-related species such as Escherichia coli and Bacillus subtilis. In spite of these obstacles, mycobacterial plasmids have been identified and DNA transformation of many mycobacterial species have now been described. The most successful method for introducing DNA into mycobacteria is electroporation. Many parameters contribute to successful transformation; these include the species/strain, the nature of the transforming DNA, the selectable marker used, the growth medium, and the  conditions for the electroporation pulse. Optimized methods for the transformation of both slow- and fast-grower are detailed here. Transformation efficiencies for different mycobacterial species and with various selectable markers are reported.

Mycobacterial pathogenic strategies remain poorly understood. The slow growth rate of most species, the impenetrable nature of the cell-wall, and the hazards of working with pathogens make mycobacteria difficult to study and are largely responsible for our poor understanding of these organisms. In this video we will demonstrate the technique of electroporation, which involves subjecting cells to a brief high electrical impulse to allow the entry of DNA. It is the most widely used method for introducing DNA into mycobacterial cells.

High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly ...

In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells

The retina and its sole output neuron, the retinal ganglion cell (RGC), comprise an excellent model in which to examine biological questions such as cell differentiation, axon guidance, retinotopic organization and synapse formation[1]. One drawback is the inability to efficiently and reliably manipulate gene expression in RGCs in vivo, especially in the otherwise accessible murine visual pathways. Transgenic mice can be used to manipulate gene expression, but this approach is often expensive, time consuming, and can produce unwanted side effects. In chick, in ovo electroporation is used to manipulate gene expression in RGCs for examining retina and RGC development. Although similar electroporation techniques have been developed in neonatal mouse pups[2], adult rats[3], and embryonic murine retinae in vitro[4], none of these strategies allow full characterization of RGC development and axon projections in vivo. To this end, we have developed two applications of electroporation, one in utero and the other ex vivo, to specifically target embryonic murine RGCs[5, 6]. 
With in utero retinal electroporation, we can misexpress or downregulate specific genes in RGCs and follow their axon projections through the visual pathways in vivo, allowing examination of guidance decisions at intermediate targets, such as the optic chiasm, or at target regions, such as the lateral geniculate nucleus. Perturbing gene expression in a subset of RGCs in an otherwise wild-type background facilitates an understanding of gene function throughout the retinal pathway. Additionally, we have developed a companion technique for analyzing RGC axon growth in vitro. We electroporate embryonic heads ex vivo, collect and incubate the whole retina, then prepare explants from these retinae several days later. Retinal explants can be used in a variety of in vitro assays in order to examine the response of electroporated RGC axons to guidance cues or other factors. In sum, this set of techniques enhances our ability to misexpress or downregulate genes in RGCs and should greatly aid studies examining RGC development and axon projections.

In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells

Here we present two techniques for manipulating gene expression in murine retinal ganglion cells (RGCs) by in utero and ex vivo electroporation. These techniques enable one to examine how alterations in gene expression affect RGC development, axon guidance, and functional properties.

The retina and its sole output neuron, the retinal ganglion cell (RGC), comprise an excellent model in which to examine biological questions such as cell ...

Using the Gene Pulser MXcell Electroporation System to Transfect Primary Cells with High Efficiency

It is becoming increasingly apparent that electroporation is the most effective way to introduce plasmid DNA or siRNA into primary cells. The Gene Pulser MXcell electroporation system and Gene Pulser electroporation buffer (Bio-Rad) were specifically developed to easily transfect nucleic acids into mammalian cells and difficult-to-transfect cells, such as primary and stem cells. We will demonstrate how to perform a simple experiment to quickly identify the best electroporation conditions. We will demonstrate how to run several samples through a range of electroporation conditions so that an experiment can be conducted at the same time as optimization is performed. We will also show how optimal conditions identified using 96-well electroporation plates can be used with standard electroporation cuvettes, facilitating the switch from electroporation plates to electroporation cuvettes while maintaining the same electroporation efficiency. In the video, we will also discuss some of the key factors that can lead to the success or failure of electroporation experiments.

This procedure shows how to use the Gene Pulser MXcell electroporation system to rapidly and easily identify the best electroporation conditions for mouse embryonic fibroblasts (MEFs) or other primary cells. Considerations for troubleshooting are also discussed in the associated video.

It is becoming increasingly apparent that electroporation is the most effective way to introduce plasmid DNA or siRNA into primary cells. The Gene Pulser ...

Reprogramming Human Somatic Cells into Induced Pluripotent Stem Cells (iPSCs) Using Retroviral Vector with GFP

Human embryonic stem cells (hESCs) are pluripotent and an invaluable cellular sources for in vitro disease modeling and regenerative medicine1. It has been previously shown that human somatic cells can be reprogrammed to pluripotency by ectopic expression of four transcription factors (Oct4, Sox2, Klf4 and Myc) and become induced pluripotent stem cells (iPSCs)2-4 . Like hESCs, human iPSCs are pluripotent and a potential source for autologous cells. Here we describe the protocol to reprogram human fibroblast cells with the four reprogramming factors cloned into GFP-containing retroviral backbone4. Using the following protocol, we generate human iPSCs in 3-4 weeks under human ESC culture condition. Human iPSC colonies closely resemble hESCs in morphology and display the loss of GFP fluorescence as a result of retroviral transgene silencing. iPSC colonies isolated mechanically under a fluorescence microscope behave in a similar fashion as hESCs. In these cells, we detect the expression of multiple pluripotency genes and surface markers.

Reprogramming Human Somatic Cells into Induced Pluripotent Stem Cells iPSCs Using Retroviral Vector with GFP

A method to generate human induced pluripotent stem cells (iPSCs) via retrovirus-mediated ectopic expression of OCT4, SOX2, KLF4 and MYC is described. A practical way to identify human iPSC colonies based on GFP expression is also discussed.

Human embryonic stem cells (hESCs) are pluripotent and an invaluable cellular sources for in vitro disease modeling and regenerative medicine1. It has ...

A Guide to Modern Quantitative Fluorescent Western Blotting with Troubleshooting Strategies

The late 1970s saw the first publicly reported use of the western blot, a technique for assessing the presence and relative abundance of specific proteins within complex biological samples. Since then, western blotting methodology has become a common component of the molecular biologists experimental repertoire. A cursory search of PubMed using the term &#8220;western blot&#8221; suggests that in excess of two hundred and twenty thousand published manuscripts have made use of this technique by the year 2014. Importantly, the last ten years have seen technical imaging advances coupled with the development of sensitive fluorescent labels which have improved sensitivity and yielded even greater ranges of linear detection. The result is a now truly Quantifiable Fluorescence based Western Blot (QFWB) that allows biologists to carry out comparative expression analysis with greater sensitivity and accuracy than ever before. Many &#8220;optimized&#8221; western blotting methodologies exist and are utilized in different laboratories. These often prove difficult to implement due to the requirement of subtle but undocumented procedural amendments. This protocol provides a comprehensive description of an established and robust QFWB method, complete with troubleshooting strategies.

The advancement of western blotting using fluorescence has allowed detection of subtle changes in protein expression enabling quantitative analyses. Here we describe a robust methodology for detection of a range of proteins across a variety of species and tissue types. A strategy to overcome common technical problems is also provided.

The late 1970s saw the first publicly reported use of the western blot, a technique for assessing the presence and relative abundance of specific proteins ...

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. 
We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisi&#230;, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.

Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing ...

Genetic Barcoding with Fluorescent Proteins for Multiplexed Applications

Fluorescent proteins, fluorescent dyes and fluorophores in general have revolutionized the field of molecular cell biology. In particular, the discovery of fluorescent proteins and their genes have enabled the engineering of protein fusions for localization, the analysis of transcriptional activation and translation of proteins of interest, or the general tracking of individual cells and cell populations. The use of fluorescent protein genes in combination with retroviral technology has further allowed the expression of these proteins in mammalian cells in a stable and reliable manner. Shown here is how one can utilize these genes to give cells within a population of cells their own biosignature. As the biosignature is achieved with retroviral technology, cells are barcoded &#180;indefinitely&#180;. As such, they can be individually tracked within a mixture of barcoded cells and utilized in more complex biological applications. The tracking of distinct populations in a mixture of cells is ideal for multiplexed applications such as discovery of drugs against a multitude of targets or the activation profile of different promoters. The protocol describes how to elegantly develop and amplify barcoded mammalian cells with distinct genetic fluorescent markers, and how to use several markers at once or one marker at different intensities. Finally, the protocol describes how the cells can be further utilized in combination with cell-based assays to increase the power of analysis through multiplexing.

Since the discovery of the green fluorescent protein gene, fluorescent proteins have impacted molecular cell biology. This protocol describes how expression of distinct fluorescent proteins through genetic engineering is used for barcoding individual cells. The procedure enables tracking distinct populations in a cell mixture, which is ideal for multiplexed applications.

Fluorescent proteins, fluorescent dyes and fluorophores in general have revolutionized the field of molecular cell biology. In particular, the discovery ...

Method for Measuring the Activity of Deubiquitinating Enzymes in Cell Lines and Tissue Samples

The ubiquitin-proteasome system has recently been implicated in various pathologies including neurodegenerative diseases and cancer. In light of this, techniques for studying the regulatory mechanism of this system are essential to elucidating the cellular and molecular processes of the aforementioned diseases. The use of hemagglutinin derived ubiquitin probes outlined in this paper serves as a valuable tool for the study of this system. This paper details a method that enables the user to perform assays that give a direct visualization of deubiquitinating enzyme activity. Deubiquitinating enzymes control proteasomal degradation and share functional homology at their active sites, which allows the user to investigate the activity of multiple enzymes in one assay. Lysates are obtained through gentle mechanical cell disruption and incubated with active site directed probes. Functional enzymes are tagged with the probes while inactive enzymes remain unbound. By running this assay, the user obtains information on both the activity and potential expression of multiple deubiquitinating enzymes in a fast and easy manner. The current method is significantly more efficient than using individual antibodies for the predicted one hundred deubiquitinating enzymes in the human cell.

The current protocol details a method for measuring the activity of functionally homologous deubiquitinating enzymes. Specialized probes covalently modify the enzyme and allow for detection. This method holds the potential to identify new therapeutic targets.

The ubiquitin-proteasome system has recently been implicated in various pathologies including neurodegenerative diseases and cancer. In light of this, ...

Comparison of Three Different Methods for Determining Cell Proliferation in Breast Cancer Cell Lines

Measuring cell proliferation can be performed by a number of different methods, each with varying levels of sensitivity, reproducibility and compatibility with high-throughput formatting. This protocol describes the use of three different methods for measuring cell proliferation in vitro including conventional hemocytometer counting chamber, a luminescence-based assay that utilizes the change in the metabolic activity of viable cells as a measure of the relative number of cells, and a multi-mode cell imager that measures cell number using a counting algorithm. Each method presents its own advantages and disadvantages for the measurement of cell proliferation, including time, cost and high-throughput compatibility. This protocol demonstrates that each method could accurately measure cell proliferation over time, and was sensitive to detect growth at differing cellular densities. Additionally, measurement of cell proliferation using a cell imager was able to provide further information such as morphology, confluence and allowed for a continual monitoring of cell proliferation over time. In conclusion, each method is capable of measuring cell proliferation, but the chosen method is user-dependent.

This protocol describes the use of three different methods for analyzing cell proliferation in breast cancer cell lines. This includes the use of conventional cell counting, luminescence-based cell viability, and cell counting through the use of a cell imager. Each offers advantages for the reproducible measurement of cell proliferation.

Measuring cell proliferation can be performed by a number of different methods, each with varying levels of sensitivity, reproducibility and compatibility ...

An Electroporation Method to Transform Rickettsia spp. with a Fluorescent Protein-Expressing Shuttle Vector in Tick Cell Lines

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