This work was supported by K12 HD068322 (SCM); R01 CA179921 and P51 OD011092 (AVM); and award 14PRE20320014 from the America Heart Association (SB).

NOTE: All procedures described in this protocol should be performed under BSL-2 conditions.
1. KSHV Stock Preparation
<ol>
	<li>Prepare TNE buffer: dissolve 292.24 mg EDTA in ddH2O, bring to 225 mL, and adjust to pH 8. Dissolve 605.7 mg Tris in ddH2O, bring to 225 mL, and adjust to pH 8. Combine EDTA and Tris solutions, add 4.38 g NaCl, adjust final volume to 500 mL, filter sterilize, and store at 4 &#176;C.</li>
	<li>Culture the KSHV-positive, EBV-negative primary effusion lymphoma cell line BCBL-1 in a humidified incubator at 37 &#176;C plus 5% CO2 in RPMI supplemented with 10% heat-inactivated fetal bovine serum (FBS) and antimicrobials to approximately 1 to 1.2 x 106 cells per mL.</li>
	<li>To induce KSHV production, split cultures 1:2 with fresh medium and add Phorbol 12-Myristate 13-Acetate (PMA) to a final concentration of 20 ng/mL and incubate cultures for 5 days. As an alternative, treat cells with sodium butyrate (NaB; 0.1 mM) for 3 days to induce lytic viral replication.</li>
	<li>To harvest viral particles, first centrifuge the culture supernatant at 300 x g for 5 min at room temperature to pellet the cells and transfer supernatants to fresh tubes.</li>
	<li>To further clarify the supernatant so as to avoid transfer of cellular debris, centrifuge at 2,500 x g for 10 min at 4 &#176;C. 
	NOTE: As an alternative to high speed centrifugation for clarification of culture supernatants, filtration through a sterile 0.45 &#181;m filter can be performed. We have observed a significant drop in titer using filtration; therefore, we routinely perform the second centrifugation step instead.</li>
	<li>Next, overlay 5 mL of 25% sucrose solution in TNE buffer with approximately 30 mL of the clarified culture supernatant in 6 ultracentrifuge tubes.</li>
	<li>Centrifuge balanced tubes at 75,000 x g for 2 h at 4 &#176;C under vacuum.</li>
	<li>Decant culture supernatant and blot the rim of each tube to remove as much supernatant and sucrose solution as possible, then resuspend the virus pellet, which may not be visible, in 150 &#181;L of TNE buffer.</li>
	<li>Pool all resuspended virus, mix well, and freeze 25 &#181;L aliquots at -80 &#176;C.</li>
</ol>
2. Transduction of EC with E6 and E7 Papillomavirus Genes
NOTE: We routinely use standard tissue culture flasks for growing primary EC. However, if unsatisfactory growth of EC are obtained then the use of commercial culture flasks should be considered.
<ol>
	<li>Culture primary human dermal microvascular or lymphatic EC in a humidified incubator at 37&#176;C plus 5% CO2 in EC growth medium (EGM; contains EC basal medium [EBM] supplemented with EGM-2 BulletKit [containing FBS, hydrocortisone, human fibroblast growth factor-B, vascular EC growth factor, insulin-like growth factor-1, ascorbic acid, human epithelial growth factor, antibiotics, and heparin]) in a T75 cell culture flask to approximately 50% confluence.</li>
	<li>For transduction, culture PA317 LXSN 16E6E7 cells24,35 in a humidified incubator at 37&#176;C plus 5% CO2 in DMEM supplemented with 10% FBS and antimicrobials in a T150 cell culture flask until they are approximately 90% confluent. Then incubate overnight in 16 mL of medium.</li>
	<li>Clarify the PA317 medium by centrifuging at 300 x g for 5 min at room temperature.</li>
	<li>Remove the EGM from the EC culture and overlay cells with 12 mL of the clarified PA317 medium and incubate for 4 h. 
	NOTE: Centrifuging 16 mL of conditioned medium at 300 x g for 5 min results in a compact pellet that is not disturbed by the careful subsequent removal of 12 mL of clarified supernatant. However, if transfer of PA317 LXSN 16E6E7 cells to primary EC cultures is a concern (the packaging cell line will quickly outgrow EC), then filtration of the conditioned supernatant through a 0.45 &#181;m filter before transfer can be performed.</li>
	<li>Replace 6 mL of the PA317 medium with fresh EGM and incubate overnight.</li>
	<li>Refeed EC with 12 mL fresh EGM and incubate a further 48 h.</li>
	<li>To sub-culture the EC, remove medium and wash with 12 mL of PBS without cations, and add 3 mL of commercial enzymatic dissociation solution (e.g., TrypLE) by and incubate at 37 &#176;C for 3 min.</li>
	<li>Transfer cell suspension to a 15 mL conical tube and centrifuge at 300 x g for 5 min at room temperature. Resuspend the resulting cell pellet in fresh EGM and divide evenly into 3 x T75 flasks with a final volume of 12 mL EGM per flask.</li>
	<li>To select for EC transduced with E6 and E7, add G418 to a final concentration of 200 &#181;g/mL for two passages, after which the transduced EC can be plated for infection or frozen in liquid nitrogen.</li>
</ol>
3. Infection of Transduced EC with KSHV
<ol>
	<li>On the day prior to infection harvest EC by enzymatic digestion using the commercial enzymatic dissociation solution as described above. Resuspend cell pellet in 1 mL of EGM. Use 5 &#181;L of cells to prepare a 1/10 dilution in Trypan blue. Count live cells using a hemocytometer and seed 2.5 x 105 live cells in 2 mL of EGM per well in 6 well plates.</li>
	<li>On the day of infection, remove EGM and wash cells with 3 mL of PBS with calcium and magnesium per well, and then add 2 mL EBM. 
	NOTE: It is essential to use EBM rather than EGM during infection, as the heparin in EGM will inhibit binding of viral particles to and subsequent infection of target EC.</li>
	<li>To infect cells with KSHV, add 5 to 20 &#181;L virus stock to each well and swirl plate to mix. For mock-infected cells add an equal volume of TNE buffer to each well.</li>
	<li>Centrifuge plates at 400 x g for 30 min at room temperature and then incubate the plates at 37 &#176;C for 90 min.</li>
	<li>If the goal of the study is to investigate early events requiring a synchronous viral infection, e.g., early events in de novo infection, remove the viral inoculum at this point. Rinse and refeed the cells with 2 mL fresh EGM. Otherwise, add 2 mL EGM and incubate cultures overnight.</li>
	<li>Refeed cells with 2 mL EGM the day after infection and then every other day.</li>
	<li>When cells are approximately 90% confluent, harvest by enzymatic digestion using commercial enzymatic dissociation solution as described above, then pool the cells from three wells into a T75, noting both the total passage number and the passage post-infection.</li>
	<li>Expand mock- and KSHV-infected EC with 1:3 splits for at least two more passages at which time cells can be used in experiments or frozen in liquid nitrogen. 
	NOTE: As with splitting of EC transduced with E6 and E7, maintenance cultures of mock- and KSHV-infected cells should be split before reaching confluence (~85 to 90%).</li>
</ol>
4. Confirming Infection with KSHV by Immunofluorescence
NOTE: Detection of the KSHV latency-associated nuclear antigen (LANA-1/ORF73) provides a reliable quantitative measure of infection. Anti-LANA antibodies are commercially available.
<ol>
	<li>The day prior to staining, plate two wells each with mock- or KSHV-infected EC on a 24-well plate at 1 x 105 cells per well in 1 mL EGM and incubate overnight.</li>
	<li>Wash cells twice with 500 &#181;L of PBS plus calcium and magniseum. Wash for 30 s on a rocking platform.</li>
	<li>In an externally-ventillated fume hood, fix cells with 500 &#181;L of 4% paraformaldehyde for 15 min at room temperature.</li>
	<li>Wash cells three times with 500 &#181;L of wash buffer (0.1% Triton X-100 plus 0.02% goat serum in PBS plus cations).</li>
	<li>Block cells with 500 &#181;L of 2% goat serum in wash buffer for 30 min at room temperature.</li>
	<li>Wash cells three times with 500 &#181;L of wash buffer.</li>
	<li>Label one well of mock- or KSHV-infected EC with 200 &#181;L of primary antibody diluted 1:100 in wash buffer for 30 min at room temperature on a rocker. Incubate the remaining two wells in 200 &#181;L of wash buffer only.</li>
	<li>Wash cells three times with 500 &#181;L of wash buffer.</li>
	<li>Label all four wells with 200 &#181;L of secondary antibody diluted 1:100 in wash buffer for 30 min at room temperature on a rocker.</li>
	<li>Wash cells three times with 500 &#181;L of wash buffer.</li>
	<li>To each well apply 20 &#181;L of mounting medium containing DAPI and a coverslip and evaluate staining using an inverted fluorescent microscope.</li>
	<li>Determine the percent infection of KSHV-infected cells by counting the number of LANA-positive nuclei in at least 200 cells. Infection with recombinant KSHV-Bac16 can also be monitored via observation of cultures for expression of GFP.</li>
</ol>

Oncogenesis is a multistep process that circumvents important safeguards within an organism36. As KS lesions exist along a spectrum of chronic inflammation to true sarcomas, elucidation of certain pathophysiological processes mediated by KSHV requires that some studies be conducted in cell culture models that support transformation9. It should be noted that loss of contact inhibition and anchorage-dependent growth, phenotypes indicative of cellular transformation, do not re...

Kaposi sarcoma (KS) is a multi-focal angioproliferative tumor affecting dermal, mucosal, and visceral sites that develops most commonly in the setting of advanced immune suppression1. Four epidemiological forms have been described: classic, an indolent form that typically affects older people of Mediterranean and Middle Eastern heritage; iatrogenic, resulting from treatment with immunosuppressive drugs following organ transplantation; epidemic, an AIDS-defining cancer; and endemic, an HIV-independent form common in children in endemic regions in Africa. With the advent of effective combination anti-retroviral drug regimens for the treatment of HIV, epidemic KS is much less commonly diagnosed in developing countries. However, the clinically aggressive endemic and epidemic forms remain among the most commonly diagnosed cancers in many African countries2,3,4. Therefore, identification of effective pathogenesis-targeted drugs for treatment of KS is a research priority.
Histologically, KS lesions are characterized by extensive but abnormal neovascularization whereby spindle cells of EC origin form discontinuous vascular networks5. These abnormal vessels (&#34;vascular slits&#34;) allow extravasation of erythrocytes, which give lesions their characteristic color. Additionally, lesions contain numerous leukocytes that characterize chronic inflammation (i.e., lymphocytes, macrophages, and plasma cells). Regression of KS lesions following immune reconstitution has been described, suggesting that KS has features of both a hyper-proliferative lesion and a true tumor6,7,8,9.
KS herpesvirus (KSHV), the causative agent of KS, was identified in 199410. Since then many in vitro cell culture models have been developed to enable pathogenesis studies, including cells explanted from tumor biopsy material and primary or telomerase-expressing EC infected with KSHV in vitro11,12,13,14,15,16,17,18. None of the currently available models fully recapitulates the KS tumor microenvironment, but all have contributed valuable knowledge to our understanding of the pathobiology of KSHV infection. Unlike the other known tumorigenic human herpesvirus Epstein-Barr virus (EBV), KSHV does not readily transform cells in culture following de novo infection19,20,21,22. However, this limitation has been overcome by transducing primary human EC of either mixed microvascular or lymphatic origin with the E6 and E7 genes from human papillomavirus type 16 prior to infection with KSHV23,24. Expression of these exogenous oncogenes dramatically increases the transforming potential of KSHV in vitro in part by providing further inhibition of the retinoblastoma protein and p5323,24. This EC transduction method has allowed multiple laboratories to identify key alterations in host cell gene expression that are induced by KSHV infection and that appear to facilitate KS cell survival and proliferation25,26,27,28,29,30,31,32. The protocols described herein are straightforward and highly reproducible, and will result in the generation of age- and passage-matched KSHV-infected EC and mock-infected controls that can be cultured for far longer than primary cells and will allow for the investigation of oncogenic mechanisms employed by KSHV. Although the protocol includes a method for production of wild type KSHV from the primary effusion lymphoma cell line BCBL-1, E6/E7-immortalized EC are also highly susceptible to infection with recombinant BACmid derived KSHV-BAC1630. Protocols for preparation of BAC16 are described elsewhere33,34.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>BCBL-1 cells</td>
			<td>NIH AIDS Reagent Program</td>
			<td>3233</td>
		</tr>
		<tr>
			<td>PA317 cells</td>
			<td>ATCC</td>
			<td>CRL-2203</td>
		</tr>
		<tr>
			<td>Neonatal dermal microvascular endothelial cells</td>
			<td>Lonza</td>
			<td>CC-2505</td>
		</tr>
		<tr>
			<td>EBM-2 Basal Medium</td>
			<td>Lonza</td>
			<td>CC-3156</td>
		</tr>
		<tr>
			<td>EGM-2 BulletKit</td>
			<td>Lonza</td>
			<td>CC-3162</td>
		</tr>
		<tr>
			<td>anti-KSHV LANA/ORF 73</td>
			<td>Advanced Biotechnologies</td>
			<td>13-210-100</td>
		</tr>
		<tr>
			<td>TrypLETMExpress, no phenol red</td>
			<td>ThermoFisher</td>
			<td>12604013</td>
		</tr>
		<tr>
			<td>RPMI</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PBS with calcium and magnesium</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

an in vitro model for studying cellular transformation by kaposi sarcoma herpesvirus

Kaposi sarcoma (KS) is an unusual tumor composed of proliferating spindle cells that is initiated by infection of endothelial cells (EC) with KSHV, and develops most often in the setting of immunosuppression. Despite decades of research, optimal treatment of KS remains poorly defined and clinical outcomes are especially unfavorable in resource-limited settings. KS lesions are driven by pathological angiogenesis, chronic inflammation, and oncogenesis, and various in vitro cell culture models have been developed to study these processes. KS arises from KSHV-infected cells of endothelial origin, so EC-lineage cells provide the most appropriate in vitro surrogates of the spindle cell precursor. However, because EC have a limited in vitro lifespan, and as the oncogenic mechanisms employed by KSHV are less efficient than those of other tumorigenic viruses, it has been difficult to assess the processes of transformation in primary or telomerase-immortalized EC. Therefore, a novel EC-based culture model was developed that readily supports transformation following infection with KSHV. Ectopic expression of the E6 and E7 genes of human papillomavirus type 16 allows for extended culture of age- and passage-matched mock- and KSHV-infected EC and supports the development of a truly transformed (i.e., tumorigenic) phenotype in infected cell cultures. This tractable and highly reproducible model of KS has facilitated the discovery of several essential signaling pathways with high potential for translation into clinical settings.

The morphology of primary EC is classically described as &#34;cobble stone&#34;, and this morphology is not altered by expression of the papillomavirus E6 and E7 genes (Figure 1A). Expression of the E6 and E7 genes alone does not induce a transformed phenotype; thus, cells are susceptible to contact inhibition and will cease dividing upon reaching confluence in culture. The cells will however proliferate and regrow to confluence upon trypsinization and replating at a lowe...

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An In Vitro Model for Studying Cellular Transformation by Kaposi Sarcoma Herpesvirus

Kaposi sarcoma (KS) is a tumor induced by infection with the oncogenic virus human herpesvirus-8/KS herpesvirus (HHV-8/KSHV). The endothelial cell culture model described here is uniquely suited for studying the mechanisms by which KSHV transforms host cells.

An In Vitro Model for Studying Cellular Transformation by Kaposi Sarcoma Herpesvirus

Kaposi sarcoma (KS) is an unusual tumor composed of proliferating spindle cells that is initiated by infection of endothelial cells (EC) with KSHV, and ...

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8.2K Views.  University of Minnesota Medical School. Kaposi sarcoma (KS) is a tumor induced by infection with the oncogenic virus human herpesvirus-8/KS herpesvirus (HHV-8/KSHV). The endothelial cell culture model described here is uniquely suited for studying the mechanisms by which KSHV transforms host cells. 

Video: An In Vitro Model for Studying Cellular Transformation by Kaposi Sarcoma Herpesvirus

Christopher Hughes:  An in vitro model for the Study of Angiogenesis (Interview)

Christopher C.W. Hughes describes the utility of his culture system for studying angiogenesis in vitro.  He explains the importance of fibroblasts that secrete a critical, yet unidentified, soluble factor that allow endothelial cells to form vessels in culture that branch, form proper lumens, and undergo anastamosis.

Christopher Hughes:  An in vitro model for the Study of Angiogenesis Interview

Christopher C.W. Hughes describes the utility of his culture system for studying angiogenesis in vitro. He explains the importance of fibroblasts that secrete a critical, yet unidentified, soluble factor that allow endothelial cells to form vessels in culture that branch, form proper lumens, and undergo anastamosis.

Christopher C.W. Hughes describes the utility of his culture system for studying angiogenesis in vitro.  He explains the importance of fibroblasts that ...

A Microfluidic Device with Groove Patterns for Studying Cellular Behavior

We describe a microfluidic device with microgrooved patterns for studying cellular behavior. This microfluidic platform consists of a top fluidic channel and a bottom microgrooved substrate. To fabricate the microgrooved channels, a top poly(dimethylsiloxane) (PDMS) mold containing the impression of the microfluidic channels was aligned and bonded to a microgrooved substrate. Using this device, mouse fibroblast cells were immobilized and patterned within microgrooved substrates (25, 50, 75, and 100 &mu;m wide). To study apoptosis in a microfluidic device, media containing hydrogen peroxide, Annexin V, and propidium iodide was perfused into the fluidic channel for 2 hours. We found that cells exposed to the oxidative stress became apoptotic. These apoptotic cells were confirmed by Annexin V that bound to phosphatidylserine at the outer leaflet of the plasma membrane during the apoptosis process. Using this microfluidic device with microgrooved patterns, the apoptosis process was observed in real-time and analyzed by using an inverted microscope containing an incubation chamber (37&deg;C, 5% CO2). Therefore, this microfluidic device incorporated with microgrooved substrates could be useful for studying the cellular behavior and performing high-throughput drug screening.

We describe a protocol for the fabrication of microfluidic devices that can enable cell capture and culture. In this approach patterned microstructures such as grooves within microfluidic channels are used to create low shear stress regions within which cell can dock.

We describe a microfluidic device with microgrooved patterns for studying cellular behavior.  This microfluidic platform consists of a top fluidic channel ...

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of the Drosophila melanogaster syncytial embryo for studying mitosis1. Drosophila has useful genetics with a sequenced genome, and it can be easily maintained and manipulated. Many mitotic mutants exist, and transgenic flies expressing functional fluorescently (e.g. GFP) - tagged mitotic proteins have been and are being generated. Targeted gene expression is possible using the GAL4/UAS system2. 

The Drosophila early embryo carries out multiple mitoses very rapidly (cell cycle duration, &asymp;10 min). It is well suited for imaging mitosis, because during cycles 10-13, nuclei divide rapidly and synchronously without intervening cytokinesis at the surface of the embryo in a single monolayer just underneath the cortex. These rapidly dividing nuclei probably use the same mitotic machinery as other cells, but they are optimized for speed; the checkpoint is generally believed to not be stringent, allowing the study of mitotic proteins whose absence would cause cell cycle arrest in cells with a strong checkpoint. Embryos expressing GFP labeled proteins or microinjected with fluorescently labeled proteins can be easily imaged to follow live dynamics (Fig. 1). In addition, embryos can be microinjected with function-blocking antibodies or inhibitors of specific proteins to study the effect of the loss or perturbation of their function3. These reagents can diffuse throughout the embryo, reaching many spindles to produce a gradient of concentration of inhibitor, which in turn results in a gradient of defects comparable to an allelic series of mutants. Ideally, if the target protein is fluorescently labeled, the gradient of inhibition can be directly visualized4. It is assumed that the strongest phenotype is comparable to the null phenotype, although it is hard to formally exclude the possibility that the antibodies may have dominant effects in rare instances, so rigorous controls and cautious interpretation must be applied. Further away from the injection site, protein function is only partially lost allowing other functions of the target protein to become evident.

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of microinjection and high resolution imaging in the Drosophila melanogaster syncytial embryo to study mitosis.

This protocol describes the use of the Drosophila melanogaster syncytial embryo  for studying mitosis1.  Drosophila has useful genetics with a sequenced ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Adenofection: A Method for Studying the Role of Molecular Chaperones in Cellular Morphodynamics by Depletion-Rescue Experiments

Cellular processes such as mitosis and cell differentiation are governed by changes in cell shape that largely rely on proper remodeling of the cell cytoskeletal structures. This involves the assembly-disassembly of higher-order macromolecular structures at a given time and location, a process that is particularly sensitive to perturbations caused by overexpression of proteins. Methods that can preserve protein homeostasis and maintain near-to-normal cellular morphology are highly desirable to determine the functional contribution of a protein of interest in a wide range of cellular processes. Transient depletion-rescue experiments based on RNA interference are powerful approaches to analyze protein functions and structural requirements. However, reintroduction of the target protein with minimum deviation from its physiological level is a real challenge. Here we describe a method termed adenofection that was developed to study the role of molecular chaperones and partners in the normal operation of dividing cells and the relationship with actin remodeling. HeLa cells were depleted of BAG3 with siRNA duplexes targeting the 3&#39;UTR region. GFP-tagged BAG3 proteins were reintroduced simultaneously into &#62;75% of the cells using recombinant adenoviruses coupled to transfection reagents. Adenofection enabled to express BAG3-GFP proteins at near physiological levels in HeLa cells depleted of BAG3, in the absence of a stress response. No effect was observed on the levels of endogenous Heat Shock Protein chaperones, the main stress-inducible regulators of protein homeostasis. Furthermore, by adding baculoviruses driving the expression of fluorescent markers at the time of cell transduction-transfection, we could dissect mitotic cell dynamics by time-lapse microscopic analyses with minimum perturbation of normal mitotic progression. Adenofection is applicable also to hard-to-infect mouse cells, and suitable for functional analyses of myoblast differentiation into myotubes. Thus adenofection provides a versatile method to perform structure-function analyses of proteins involved in sensitive biological processes that rely on higher-order cytoskeletal dynamics.

We describe a method for depletion-rescue experiments that preserves cellular integrity and protein homeostasis. Adenofection enables functional analyses of proteins within biological processes that rely on finely tuned actin-based dynamics, such as mitotic cell division and myogenesis, at the single-cell level.

Cellular processes such as mitosis and cell differentiation are governed by changes in cell shape that largely rely on proper remodeling of the cell ...

An Efficient In Vitro Transposition Method by a Transcriptionally Regulated Sleeping Beauty System Packaged into an Integration Defective Lentiviral Vector

The Sleeping Beauty (SB) transposon is a non-viral integrating system with proven efficacy for gene transfer and functional genomics. To optimize the SB transposon machinery, a transcriptionally regulated hyperactive transposase (SB100X) and T2-based transposon are employed. Typically, the transposase and transposon are provided transiently by plasmid transfection and SB100X expression is driven by a constitutive promoter. Here, we describe an efficient method to deliver the SB components to human cells that are resistant to several physical and chemical transfection methods, to control SB100X expression and stably integrate a gene of interest (GOI) through a &#34;cut and paste&#34; SB mechanism. The expression of hyperactive transposase is tightly controlled by the Tet-ON system, widely used to control gene expression since 1992. The gene of interest is flanked by inverted repeats (IR) of the T2 transposon. Both SB components are packaged in integration defective lentiviral vectors transiently produced in HEK293T cells. Human cells, either cell lines or primary cells from human tissue, are in vitro transiently transduced with viral vectors. Upon addition of doxycycline (dox, tetracycline analog) into the culture medium, a fine-tuning of transposase expression is measured and results in a long-lasting integration of the gene of interest in the genome of the treated cells. This method is efficient and applicable to the cell line (e.g., HeLa cells) and primary cells (e.g., human primary keratinocytes), and thus represents a valuable tool for genetic engineering and therapeutic gene transfer.

An Efficient In Vitro Transposition Method by a Transcriptionally Regulated Sleeping Beauty System Packaged into an Integration Defective Lentiviral Vector

This protocol describes a method to achieve stable integration of a gene of interest into the human genome by the transcriptionally regulated Sleeping Beauty system. Preparation of the integration of defective lentiviral vectors, the in vitro transduction of human cells, and the molecular assay on transduced cells are reported.

The Sleeping Beauty (SB) transposon is a non-viral integrating system with proven efficacy for gene transfer and functional genomics. To optimize the SB ...

Quantitative Approaches for Studying Cellular Structures and Organelle Morphology in Caenorhabditis elegans

Defining the cellular mechanisms underlying disease is essential for the development of novel therapeutics. A strategy frequently used to unravel these mechanisms is to introduce mutations in candidate genes and qualitatively describe changes in the morphology of tissues and cellular organelles. However, qualitative descriptions may not capture subtle phenotypic differences, might misrepresent phenotypic variations across individuals in a population, and are frequently assessed subjectively. Here, quantitative approaches are described to study the morphology of tissues and organelles in the nematode Caenorhabditis elegans using laser scanning confocal microscopy combined with commercially available bio-image processing software. A quantitative analysis of phenotypes affecting synapse integrity (size and integrated fluorescence levels), muscle development (muscle cell size and myosin filament length), and mitochondrial morphology (circularity and size) was performed to understand the effects of genetic mutations on these cellular structures. These quantitative approaches are not limited to the applications described here, as they could readily be used to quantitatively assess the morphology of other tissues and organelles in the nematode, as well as in other model organisms.

Quantitative Approaches for Studying Cellular Structures and Organelle Morphology in Caenorhabditis elegans

This study outlines quantitative measurements of synaptic size and localization, muscle morphology, and mitochondrial shape in C. elegans using freely available image processing tools. This approach allows future studies in&#160;C. elegans&#160;to quantitatively compare the extent of tissue and organelle structural changes as a result of genetic mutations.

Defining the cellular mechanisms underlying disease is essential for the development of novel therapeutics. A strategy frequently used to unravel these ...

In Vitro Analysis of E3 Ubiquitin Ligase Function

The covalent attachment of ubiquitin (Ub) to internal lysine residue(s) of a substrate protein, a process termed ubiquitylation, represents one of the most important post-translational modifications in eukaryotic organisms. Ubiquitylation is mediated by a sequential cascade of three enzyme classes including ubiquitin-activating enzymes (E1 enzymes), ubiquitin-conjugating enzymes (E2 enzymes), and ubiquitin ligases (E3 enzymes), and sometimes, ubiquitin-chain elongation factors (E4 enzymes). Here, in vitro protocols for ubiquitylation assays are provided, which allow the assessment of E3 ubiquitin ligase activity, the cooperation between E2-E3 pairs, and substrate selection. Cooperating E2-E3 pairs can be screened by monitoring the generation of free poly-ubiquitin chains and/or auto-ubiquitylation of the E3 ligase. Substrate ubiquitylation is defined by selective binding of the E3 ligase and can be detected by western blotting of the in vitro reaction. Furthermore, an E2~Ub discharge assay is described, which is a useful tool for the direct assessment of functional E2-E3 cooperation. Here, the E3-dependent transfer of ubiquitin is followed from the corresponding E2 enzyme onto free lysine amino acids (mimicking substrate ubiquitylation) or internal lysines of the E3 ligase itself (auto-ubiquitylation). In conclusion, three different in vitro protocols are provided that are fast and easy to perform to address E3 ligase catalytic functionality.

In Vitro Analysis of E3 Ubiquitin Ligase Function

The present study provides detailed in vitro ubiquitylation assay protocols for the analysis of E3 ubiquitin ligase catalytic activity. Recombinant proteins were expressed using prokaryotic systems such as Escherichia coli culture.

The covalent attachment of ubiquitin (Ub) to internal lysine residue(s) of a substrate protein, a process termed ubiquitylation, represents one of the ...

Investigating Zinc Transport Mechanism in Mammalian Cells with FluoZin-3 Assay

Characterizing Mammalian Zinc Transporters Using an In Vitro Zinc Transport Assay

Transition metals such as Zn2+ ions must be tightly regulated due to their cellular toxicity. Previously, the activity of Zn2+ transporters was measured indirectly by determining the expression level of the transporter under different concentrations of Zn2+. This was done by utilizing immunohistochemistry, measuring mRNA in the tissue, or determining the cellular Zn2+ levels. With the development of intracellular Zn2+ sensors, the activities of zinc transporters are currently primarily determined by correlating changes in intracellular Zn2+, detected using fluorescent probes, with the expression of the Zn2+ transporters. However, even today, only a few labs monitor dynamic changes in intracellular Zn2+ and use it to measure the activity of zinc transporters directly. Part of the problem is that out of the 10 zinc transporters of the ZnT family, except for ZnT10 (transports manganese), only zinc transporter 1 (ZnT1) is localized at the plasma membrane. Therefore, linking the transport activity to changes in the intracellular Zn2+ concentration is hard. This article describes a direct way to determine the zinc transport kinetics using an assay based on a zinc-specific fluorescent dye, FluoZin-3. This dye is loaded into mammalian cells in its ester form and then trapped in the cytosol due to cellular di-esterase activity. The cells are loaded with Zn2+ by utilizing the Zn2+ ionophore pyrithione. The ZnT1 activity is assessed from the linear part of the reduction in fluorescence following the cell washout. The fluorescence measured at an excitation of 470 nm and emission of 520 nm is proportional to the free intracellular Zn2+. Selecting the cells expressing ZnT1 tagged with the mCherry fluorophore allows for monitoring only the cells expressing the transporter. This assay is used to investigate the contribution of different domains of ZnT1 protein to the transport mechanism of human ZnT1, a eukaryotic transmembrane protein that extrudes excess zinc from the cell.

Author Spotlight: Exploring Cellular Zinc Regulation Through ZnT1 Functionality 

Zinc transport has proven challenging to measure due to the weak causal links to protein function and the low temporal resolution. This protocol describes a method for monitoring, with high temporal resolution, Zn2+ extrusion from living cells by utilizing a Zn2+ sensitive fluorescent dye, thus providing a direct measure of Zn2+ efflux.

Transition metals such as Zn2+ ions must be tightly regulated due to their cellular toxicity. Previously, the activity of Zn2+ transporters was measured ...