Supported by grants from the Department of Defense (W81XWH-09-1-0127), the NIH (2-R01-CA089569), the Elsa U. Pardee Foundation, a Marsha Rivkin Center for Ovarian Cancer Research Pilot Study Award, and generous philanthropic support from Section of Urology and Section of Research in the Department of Surgery, University of Chicago.

All mice were housed, maintained and euthanized according to Institutional Animal Care and Use Committee (IACUC) guidelines and under the supervision of the University of Chicago Animal Resource Center.
1. Preparing Animals for Experimental Studies
<ol>
	<li>Allow animals to acclimate to new housing and environment, to recover from potential physiologic effects of transport and handling. 
	Note: The following technique is applicable to all commercially available strains of mice. The number of animals to be used for an experiment is dependent on the study design and should be done with consultation from a statistician.</li>
</ol>
2. Identification and Isolation of Peritoneal Fat Depots.
<ol>
	<li>Euthanize animals via CO2 overdose&#160;and a secondary physical method to confirm death.&#160;</li>
	<li>As harvesting peritoneal fat depots constitutes internal organ removal (secondary method), make a midline incision close to the inguinal papillae through the peritoneal wall to expose the internal organs (Figure 1A).</li>
	<li>For the Omental Fat (OM), expose the omental/pancreatic complex by extending the spleen from the peritoneal cavity with forceps (Figure 1C). Release omentum from the pancreas and spleen by closely trimming all tissue connections. Ensure that any remaining pancreatic tissue is excised 4.</li>
	<li>For the Gonadal Fat (GF), use forceps to lift the gonadal fat surrounding the ovaries and excise by cutting tissue connections (Figure 1A upper right) 4.</li>
	<li>For the Uterine Fat (UF), use forceps to lift the uterine fat surrounding the uterine horns and excise by cutting tissue connections (Figure 1A lower right) 4.</li>
	<li>For the Mesenteric Fat (MF), cut the junction between the small intestine and the pylorus. Use forceps to firmly grip the free end of the small intestine, and slowly &#8220;peel&#8221; it away from the mesenteric fat. Release the mesenteric fat from the mesenteric root using dissecting scissors (Figure 1A lower left) 4.</li>
	<li>For the Splenoportal Fat (SF), lift the distal end of the spleen using forceps to expose the thin fatty band of tissue connecting the hilum of the spleen to the pancreas. Excise the splenoportal fat by first releasing it from the pancreas, and then carefully dissecting it from the spleen (Figure 1B) 4.</li>
</ol>
3. Milky Spot Identification Using Giemsa Staining
<ol>
	<li>Excise omenta (refer to step 2.3) from C57BL/6 mice and fix in 5% neutral buffered formalin at 4 &#176;C for 16 hr&#160;(O/N).</li>
	<li>For paraffin embedding the fixed tissue, choose a mold that is appropriate for the size of the tissue. Pour molten paraffin from the paraffin reservoir to fill the mold. Open the cassette and using warm forceps, transfer the tissue into the mold.
	<ol>
		<li>Carefully orient the tissues during paraffin embedding to produce longitudinal sections. Place the labeled tissue cassette over the molding and press firmly in place. Allow the mold to cool for at least 30 min.</li>
	</ol>
	</li>
	<li>Cut serial sections (4 &#181;m thick) from the formalin-fixed paraffin embedded (FFPE) tissue block. Place a precleaned slide under the sections as the ribbon of tissue comes off the paraffin block.</li>
	<li>Serially section the whole omentum; collect and stain every third section for Giemsa stain (refer to 3.6). Allow the glass slide with the sections to air dry, preferably O/N at RT. Dried slides are ready for fixation. Stain the first section for hematoxylin and eosin for comparisons to Giemsa stained slides.</li>
	<li>Deparaffinize the slides by two sequential 5 min incubations in xylenes. Rehydrate slides by two sequential incubations in the following: 100% ethanol, 95% ethanol, and finally tap water.
	<ol>
		<li>Take adequate precautions to avoid the slides from drying. Perform all steps at RT.</li>
	</ol>
	</li>
	<li>Completely immerse the slides in a Coplin jar containing 5% Giemsa solution (w/v prepared in tap water) and incubate for 4 min&#160;at RT. Rinse slides in tap water for 60 sec, air-dry and mount with coverslip using mounting media.</li>
	<li>Image the stained slides using a Whole Slide Scanner and process with manufacturer software. Quantify the milky spot volume in the Giemsa-stained omental sections using ImageJ software4.</li>
</ol>
4. Propagation and Preparation of Ovarian Cancer Cells for In vivo and Ex vivo&#160;Studies
<ol>
	<li>Pre-warm 15 ml of cell growth medium to 37 &#176;C in a heat bath. Prepare an appropriately sized tissue culture flask (e.g., 75 cm2 surface area) by labeling with name of cell line, passage number, date and person who prepared the frozen stock, and the date and person who is thawing/plating the cells. 
	Note: In studies of metastasis such detailed information is crucial, as the date of freeze down and passage number can affect the metastatic phenotype.</li>
	<li>Using the sterile environment of the biological safety hood, pipette 10 ml of medium into the T-75 culture flask. Remove a vial of cells from the liquid nitrogen system and check the label to confirm the cell type. Thaw only one vial of cells at a time by warming in 37 &#176;C heat bath.</li>
	<li>Using sterile technique, pipette 1x106 thawed cells into the culture flask. Place the flask in the incubator to allow cells to attach. Gently rock the flask back and forth to form a uniform cell suspension.
	<ol>
		<li>Place flask in tissue culture incubator and maintain at 37 &#176;C in 5% CO2 environment. Allow cells to adhere O/N.</li>
	</ol>
	</li>
	<li>Aspirate media, and replace with fresh pre-warmed growth media. Place the flask in the incubator and allow cells to grow. Monitor cell growth daily by visual inspection every 2-3 days using an inverted microscope. Use standard cell culture techniques to passage cells upon reaching 70-80% confluence. 
	Note: For experimental assays harvest the cells at 70% confluence to ensure that cells are actively proliferating.</li>
	<li>Aspirate the culture medium and rinse with 10 ml of sterile phosphate buffered saline (PBS). Aspirate PBS and add 0.25% trypsin/EDTA (2 to 3 ml for a 75 cm2 surface area) and incubate 5 min at 37 &#186;C. Visually confirm that the cells have detached and add an equal volume of growth medium, and gently pipette cells up and down to obtain a single cell suspension. 
	Note: It is critical that the growth medium contain serum as it inactivates trypsin. Prepare at least 2- to 4-fold more cells than the number required for a given experiment to compensate for loss of cells during subsequent steps</li>
	<li>Determine the percentage of viable cells in the suspension via trypan blue exclusion assay using a hemacytometer or an automated cell counter. 
	Note: Here, only use cell preparations in which &#8805; 96% of cells are viable.</li>
	<li>Transfer the cell suspension from the flask to a 50 ml conical tube. Pellet cells by centrifuging 5 min at 1,100 x g, at 4 oC.</li>
	<li>Aspirate the supernatant and resuspend the cell pellet in PBS to a concentration of 2x106 cells per ml.</li>
</ol>
5. Intraperitoneal Injection of Cells
Note: In our experiments, mice are handled in a laminar airflow or biosafety cabinet within our barrier facility in order to limit the risk of pathogen exposure,&#160; In order to demonstrate this technique clearly, the procedures in the accompanying video were conducted in a laboratory approved for animal work.&#160;This particular technique does not require the animals to be under anesthesia. Under approved protocols, perform this technique on live animals. Use appropriate strains of mice for this study. For example: use immunocompromised, athymic nude mice for the study of human ovarian cancer cell (SKOV3ip.1, and HeyA8) colonization; use immunocompetent C57BL/6 mice for study of mouse ovarian cancer cells (ID8) colonization.

<ol>
	<li>Load 500 &#181;l of the single cell suspension into the syringe and put in the sterile capped 25 G needle. This reduces cell shearing prior to injection.</li>
	<li>Pick the mouse up by the scruff of the neck and hold the tail using the palm and forefinger and fix the left hind leg between the ring and little finger (when the mouse is restrained with the left hand). 
	Note: To avoid traumatizing abdominal organs, restrain the mouse well so that it cannot move during the injection.</li>
	<li>Imagine a line across abdomen just above the knees, and locate a point on the animal&#8217;s right side and close to the midline. The point of entry is cranial to and slightly medial of the last nipple. 
	Note: Inserting the needle on the mouse&#8217;s right side avoids the cecum and reduces the risk of puncturing the intestines11.</li>
	<li>Insert the needle at the lower lateral region of the mice&#8217;s abdomen to a depth of approximately 0.5 cm. Pull back on the plunger to confirm that the needle has not penetrated a blood vessel or other peritoneal organs.
	<ol>
		<li>If any fluid is aspirated discard the syringe (and sample)11. A greenish-brown or yellow aspirate indicates needle penetration into the intestines or bladder, respectively.</li>
	</ol>
	</li>
	<li>Inject the sample using slow, steady pressure. Withdraw the needle and return the mouse to its cage. Do not recap syringe before disposal in sharps container.</li>
	<li>Sacrifice mice via CO2 asphyxiation and vital organ removal at specific time points post injection.</li>
</ol>
6. Milky spot colonization In vivo
<ol>
	<li>Quantitation of cancer cell localization to milky spots using flow cytometry
	<ol>
		<li>Isolate peritoneal fat depots from mice (as described in step 2.3 - 2.7) injected with fluorescently tagged cancer cells and immediate place tissues in ice-cold PBS.
		<ol>
			<li>For this particular technique, weight-match all adipose fat tissues to omentum. Transfer tissues to separate 5 ml tubes containing 1.5 ml of serum-free DMEM and 0.1% (w/v) bovine serum albumin. Pool tissues harvested from three independent mice to ensure a sufficient yield of cells for flow cytometry.</li>
		</ol>
		</li>
		<li>Mince tissues using surgical scissors and add 1.5 ml of serum-free DMEM containing 0.4% (w/v) collagenase I. Incubate the tissue suspension at 37 &#176;C for 30 min&#160;with rotational mixing.
		<ol>
			<li>As an optional step, further disassociate tissue by mastication. In this case, transfer the tissue-collagenase suspension to a microstomacher bag, masticate for 10 min on low, rotating the orientation of the bags after 5 min.</li>
		</ol>
		</li>
		<li>Filter samples through a nylon mesh filter (60 &#181;m pore) to remove larger debris. Collect cells via centrifugation at 250 x g for 5 min&#160;at 4 &#176;C and discard the supernatant fraction.</li>
		<li>Resuspend the cell pellet in 100 &#181;l of PBS combined with 900 &#181;l of ACK lysis buffer and incubate at RT for 1 min. Centrifuge cells at 250 x g for 5 min&#160;and discard supernatant.
		<ol>
			<li>Resuspend the cell pellet in 250 &#181;l of ice-cold PBS. Filter the samples through a 60 &#181;m pore nylon mesh to ensure single cell suspension. Rinse the filter with 250 &#181;l of ice-cold PBS.</li>
		</ol>
		</li>
		<li>Quantitate the number of fluorescently tagged cells in the population using a flow cytometer equipped with a 561-nm yellow-green laser and a 585 nm/15 nm bandpass filter. Set the gate for tdTomato-positive cells based on analysis of parental cells and/or appropriate negative controls 4.</li>
	</ol>
	</li>
	<li>Quantitation of microscopic and macroscopic metastases
	<ol>
		<li>At the desired experimental endpoint(s), isolate, and immediately place tissues in the appropriate fixation media. Fix larger samples, such as intact gonadal, uterine, and mesenteric fat depots in 10% neutral buffered formalin for 48 hr&#160;at 4 &#176;C. Fix smaller samples (omental and splenoportal fat, as well as tissue equivalents of uterine and mesenteric fat) in 5% neutral buffered formalin at 4 &#176;C for 2-16 hr.
		<ol>
			<li>Alternatively, snap freeze tissues by placing them in a freezing medium such as OCT and drop into an appropriate amount of liquid nitrogen.</li>
		</ol>
		</li>
		<li>Transfer fixed tissues to 70% ethanol and store at 4 &#176;C until embedding in paraffin. Embed and process tissue as described in 3.2 and 3.3.
		<ol>
			<li>Use the H&#38;E stained section to evaluate tissues for the presence or absence of microscopic metastases (i.e., clusters of 50 cells). Confirm the presence of microscopic metastases by a secondary method such as an IHC stain for cytokeratin 8/18 to detect ovarian cancer cells.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
7. Milky Spot Colonization Ex Vivo
<ol>
	<li>Establishment of basic ex vivo omental organ culture conditions
	<ol>
		<li>Place each omentum into a single culture plate insert and place within one well of a twelve-well tissue culture plate containing 500 &#181;l of DME/F12 media containing 20% FCS. Maintain organ cultures at 37 &#176;C in a 5% CO2 environment for 24 to 48 hr&#160;and/or additional time points. Use three independent omenta (triplicate samples) for each condition (e.g., media type/time point).</li>
		<li>To confirm the integrity of tissues at the experimental endpoints, fix and process samples as described in Step 3. Counting healthy versus necrotic adipocytes on H&#38;E sections can assess tissue integrity. A minimum of ~120 cells from 5 biological samples is required to formulate a percentage of live tissue present10.</li>
		<li>To measure tissue function, place omenta (n = 3) in separate wells of a 24-well plate containing 500&#160;&#181;l of DME/F12 media with 20% FCS. Allow omenta to condition the media at 37 &#176;C in a 5% CO2 environment for 24 hr, and then remove 250 &#181;l for use in an IL-6 ELISA assay10.</li>
	</ol>
	</li>
	<li>Ex vivo co-culture of the mouse omentum with SKOV3ip.1-GFP cells
	<ol>
		<li>Grow and prepare fluorescently tagged cells as described in section 4 and resuspend at a concentration of 2&#160;&#215;&#160;106 cells per ml.</li>
		<li>Apply ~6 &#181;l of the tissue adhesive to the membrane of the culture insert and allow it to air dry. Wash the membrane twice with sterile water to remove any excessive adhesive. Air dry the membranes under a laminar hood.</li>
		<li>Carefully excise the omenta and attach it to the adhesive-coated membrane using sterile forceps. Allow the tissue to adhere to the membrane for 1&#160;min prior to adding media. Add 500&#160;&#181;l of the cell suspension on top of each omentum in each culture insert. Fill the area around the transwell chamber with 2.5&#160;ml of DME/F-12 10.</li>
		<li>Incubate omenta with cell suspension for 6&#160;hr at 37 &#176;C in a 5% CO2 environment. Carefully remove and wash the omenta with&#160;~10&#160;ml of PBS. Visualize fluorescent cancer cell foci using an appropriate fluorescent imaging system 10.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Development of therapies to target disseminated cells requires a mechanistic understanding of metastatic colonization, the critical first step in the development of peritoneal disease. To address these issues we report approaches that can be used to discern how the omentum&#8217;s unique tissue composition and architecture promote ovarian cancer metastatic colonization. Distinguishing features of our approach are: 1) the focus on early events in the colonization process; 2) comparison of milky spot-containing an...

Unlike most solid tumors, metastases from high-grade serous ovarian cancer (HGSC) are limited to the peritoneal cavity 1. Thus, effective peritoneal therapies could potentially control or eradicate HGSC. Currently, a standard therapeutic approach is surgical cytoreduction combined with chemotherapy 1-3. Unfortunately, the vast majority of patients experience and succumb to complications of disease recurrence. These dismal statistics show the need for improved understanding of metastatic colonization, the process by which cancer cells localize to, utilize, and proliferate within host tissues to form metastases 2.
The omentum is a preferred early site of HGSC metastasis 4-7. Unlike other peritoneal adipose, omental adipose tissue contains unusual immune structures known as milky spots, which contain B, T, and NK cells and macrophages, which play key roles in peritoneal homeostasis8,9. In addition to their physiologic functions, we found that milky spots play an active role in ovarian cancer metastatic colonization 4. In experimental metastasis assays, SKOV3ip.1, CaOV3, and HeyA8 (human) and ID8 (murine; C57BL/6) ovarian cancer cells rapidly home to milky spots, suggesting that the cells are moving toward a secreted chemotactic factor. Interestingly, cancer cells do not colonize peritoneal adipose lacking milky spots (i.e., gonadal and uterine fat) 4.
In order to identify mechanisms regulating milky spot colonization, we have optimized xenograft models that enable the interrogation of cellular and molecular events over the time course of metastatic colonization 4. A specific advantage of the approaches described herein is its emphasis on tissue architecture and function, which enables users to test hypotheses in fully integrated in vivo and ex vivo models of metastatic colonization 4,10. By comparing cancer cell localization and growth in peritoneal fat depots which either contain or lack milky spots, investigators can test the relative contribution(s) of adipocytes and cells within milky spots to the lodging and progressive growth of ovarian cancer cells in physiologically relevant tissues.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Dissection Tools</td>
			<td>Fine Science Tools</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Geimsa</td>
			<td>Fluka/Sigma Aldrich</td>
			<td>48900</td>
			<td></td>
		</tr>
		<tr>
			<td>5% Formalin</td>
			<td>Sigma</td>
			<td>HT501320</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Corning</td>
			<td>10-013-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin</td>
			<td>Gibco</td>
			<td>25200-056</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Corning</td>
			<td>21-040-CV</td>
			<td>Without calcium and Magnesium</td>
		</tr>
		<tr>
			<td>26 gauge needle</td>
			<td>BD</td>
			<td>329652</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma</td>
			<td>A7906</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase</td>
			<td>Worthington</td>
			<td>LS004196</td>
			<td></td>
		</tr>
		<tr>
			<td>Stomacher</td>
			<td>Seward Labsystems</td>
			<td>Stomacher 80 Biomaster</td>
			<td></td>
		</tr>
		<tr>
			<td>Microstomacher bag</td>
			<td>Stomacher Lab Systems</td>
			<td>BA6040/Micro</td>
			<td></td>
		</tr>
		<tr>
			<td>ACK Lysis Buffer</td>
			<td>Gibco</td>
			<td>A10492-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Millicell culture plate insert</td>
			<td>Millipore</td>
			<td>PICM01250</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell-Tak</td>
			<td>BD</td>
			<td>354240</td>
			<td></td>
		</tr>
	</tbody>
</table>

in vivo and ex vivo approaches to study ovarian cancer metastatic colonization of milky spot structures in peritoneal adipose

High-grade serous ovarian cancer (HGSC), the cause of widespread peritoneal metastases, continues to have an extremely poor prognosis; fewer than 30% of women are alive 5 years after diagnosis. The omentum is a preferred site of HGSC metastasis formation. Despite the clinical importance of this microenvironment, the contribution of omental adipose tissue to ovarian cancer progression remains understudied. Omental adipose is unusual in that it contains structures known as milky spots, which are comprised of B, T, and NK cells, macrophages, and progenitor cells surrounding dense nests of vasculature. Milky spots play a key role in the physiologic functions of the omentum, which are required for peritoneal homeostasis. We have shown that milky spots also promote ovarian cancer metastatic colonization of peritoneal adipose, a key step in the development of peritoneal metastases. Here we describe the approaches we developed to evaluate and quantify milky spots in peritoneal adipose and study their functional contribution to ovarian cancer cell metastatic colonization of omental tissues both in vivo and ex vivo. These approaches are generalizable to additional mouse models and cell lines, thus enabling the study of ovarian cancer metastasis formation from initial localization of cells to milky spot structures to the development of widespread peritoneal metastases.

Identification and Histologic Examination of Peritoneal Fat Depots
Gross anatomic dissection allows identification of four of the five primary sources of peritoneal fat (Figure 1A). Moving clockwise from the top center are: the omentum (OM; outlined) located over the stomach and spleen, the gonadal fat (GF) surrounding the left ovary (ov), the uterine fat (UF) attached to the uterine horns (uh) and the mesentery (MY) attached to the small intestine (si). The o...

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In Vivo and Ex Vivo Approaches to Study Ovarian Cancer Metastatic Colonization of Milky Spot Structures in Peritoneal Adipose

We outline a protocol that implements both in vivo and ex vivo approaches to study ovarian cancer colonization of peritoneal adipose tissues, particularly the omentum. Furthermore, we present a protocol to quantitate and analyze immune cell-structures in the omentum known as milky spots, which promote metastases of peritoneal adipose.

In Vivo and Ex Vivo Approaches to Study Ovarian Cancer Metastatic Colonization of Milky Spot Structures in Peritoneal Adipose

High-grade serous ovarian cancer (HGSC), the cause of widespread peritoneal metastases, continues to have an extremely poor prognosis; fewer than 30% of ...

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13.0K ビュー.  The University of Chicago. We outline a protocol that implements both in vivo and ex vivo approaches to study ovarian cancer colonization of peritoneal adipose tissues, particularly the omentum. Furthermore, we present a protocol to quantitate and analyze immune cell-structures in the omentum known as milky spots, which promote metastases of peritoneal adipose.

ビデオ: In Vivo and Ex Vivo Approaches to Study Ovarian Cancer Metastatic Colonization of Milky Spot Structures in Peritoneal Adipose

Ex Vivo Infection of Live Tissue with Oncolytic Viruses

Oncolytic Viruses (OVs) are novel therapeutics that selectively replicate in and kill tumor cells1. Several clinical trials evaluating the effectiveness of a variety of oncolytic platforms including HSV, Reovirus, and Vaccinia OVs as treatment for cancer are currently underway2-5. One key characteristic of oncolytic viruses is that they can be genetically modified to express reporter transgenes which makes it possible to visualize the infection of tissues by microscopy or bio-luminescence imaging6,7. This offers a unique advantage since it is possible to infect tissues from patients ex vivo prior to therapy in order to ascertain the likelihood of successful oncolytic virotherapy8. To this end, it is critical to appropriately sample tissue to compensate for tissue heterogeneity and assess tissue viability, particularly prior to infection9. It is also important to follow viral replication using reporter transgenes if expressed by the oncolytic platform as well as by direct titration of tissues following homogenization in order to discriminate between abortive and productive infection. The object of this protocol is to address these issues and herein describes 1. The sampling and preparation of tumor tissue for cell culture 2. The assessment of tissue viability using the metabolic dye alamar blue 3. Ex vivo infection of cultured tissues with vaccinia virus expressing either GFP or firefly luciferase 4. Detection of transgene expression by fluorescence microscopy or using an In Vivo Imaging System (IVIS) 5. Quantification of virus by plaque assay. This comprehensive method presents several advantages including ease of tissue processing, compensation for tissue heterogeneity, control of tissue viability, and discrimination between abortive infection and bone fide viral replication.

Ex Vivo Infection of Live Tissue with Oncolytic Viruses

Oncolytic viruses are promising for cancer therapeutics. The ability to ascertain the infectability of live tissue specimens obtained from patients prior to treatment is a unique advantage of this therapeutic approach. This protocol describes how to process tissues for ex vivo infection with oncolytic virus and subsequent viral quantification.

Oncolytic Viruses (OVs) are novel therapeutics that selectively replicate in and kill tumor cells1. Several clinical trials evaluating the effectiveness ...

An In Vitro System to Study Tumor Dormancy and the Switch to Metastatic Growth

Recurrence of breast cancer often follows a long latent period in which there are no signs of cancer, and metastases may not become clinically apparent until many years after removal of the primary tumor and adjuvant therapy. A likely explanation of this phenomenon is that tumor cells have seeded metastatic sites, are resistant to conventional therapies, and remain dormant for long periods of time 1-4.
The existence of dormant cancer cells at secondary sites has been described previously as quiescent solitary cells that neither proliferate nor undergo apoptosis 5-7. Moreover, these solitary cells has been shown to disseminate from the primary tumor at an early stage of disease progression 8-10 and reside growth-arrested in the patients' bone marrow, blood and lymph nodes 1,4,11. Therefore, understanding mechanisms that regulate dormancy or the switch to a proliferative state is critical for discovering novel targets and interventions to prevent disease recurrence. However, unraveling the mechanisms regulating the switch from tumor dormancy to metastatic growth has been hampered by the lack of available model systems.
in vivo and ex vivo model systems to study metastatic progression of tumor cells have been described previously 1,12-14. However these model systems have not provided in real time and in a high throughput manner mechanistic insights into what triggers the emergence of solitary dormant tumor cells to proliferate as metastatic disease. We have recently developed a 3D in vitro system to model the in vivo growth characteristics of cells that exhibit either dormant (D2.OR, MCF7, K7M2-AS.46) or proliferative (D2A1, MDA-MB-231, K7M2) metastatic behavior in vivo . We demonstrated that tumor cells that exhibit dormancy in vivo at a metastatic site remain quiescent when cultured in a 3-dimension (3D) basement membrane extract (BME), whereas cells highly metastatic in vivo readily proliferate in 3D culture after variable, but relatively short periods of quiescence. Importantly by utilizing the 3D in vitro model system we demonstrated for the first time that the ECM composition plays an important role in regulating whether dormant tumor cells will switch to a proliferative state and have confirmed this in in vivo studies15-17. Hence, the model system described in this report provides an in vitro method to model tumor dormancy and study the transition to proliferative growth induced by the microenvironment.

An In Vitro System to Study Tumor Dormancy and the Switch to Metastatic Growth

A modified 3-D in vitro system is presented in which growth characteristics of several tumor cell lines in reconstituted basement membrane correlate with the dormant or proliferative behavior of the tumor cells at a metastatic secondary site in vivo.

Recurrence of breast cancer often follows a long latent period in which there are no signs of cancer, and metastases may not become clinically apparent ...

A Novel High-resolution In vivo Imaging Technique to Study the Dynamic Response of Intracranial Structures to Tumor Growth and Therapeutics

We have successfully integrated previously established Intracranial window (ICW) technology 1-4 with intravital 2-photon confocal microscopy to develop a novel platform that allows for direct long-term visualization of tissue structure changes intracranially. Imaging at a single cell resolution in a real-time fashion provides supplementary dynamic information beyond that provided by standard end-point histological analysis, which looks solely at 'snap-shot' cross sections of tissue.
Establishing this intravital imaging technique in fluorescent chimeric mice, we are able to image four fluorescent channels simultaneously. By incorporating fluorescently labeled cells, such as GFP+ bone marrow, it is possible to track the fate of these cells studying their long-term migration, integration and differentiation within tissue. Further integration of a secondary reporter cell, such as an mCherry glioma tumor line, allows for characterization of cell:cell interactions. Structural changes in the tissue microenvironment can be highlighted through the addition of intra-vital dyes and antibodies, for example CD31 tagged antibodies and Dextran molecules.
Moreover, we describe the combination of our ICW imaging model with a small animal micro-irradiator that provides stereotactic irradiation, creating a platform through which the dynamic tissue changes that occur following the administration of ionizing irradiation can be assessed.
Current limitations of our model include penetrance of the microscope, which is limited to a depth of up to 900 &mu;m from the sub cortical surface, limiting imaging to the dorsal axis of the brain. The presence of the skull bone makes the ICW a more challenging technical procedure, compared to the more established and utilized chamber models currently used to study mammary tissue and fat pads 5-7. In addition, the ICW provides many challenges when optimizing the imaging.

A Novel High-resolution In vivo Imaging Technique to Study the Dynamic Response of Intracranial Structures to Tumor Growth and Therapeutics

We describe a novel in vivo imaging technique that couples fluorescent chimeric mice with intracranial windows and high-resolution 2-photon microscopy. This imaging platform aids studies of dynamic changes in brain tissue and microvasculature, at a single-cell level, following pathological insults and is adaptable to assess intracranial drug delivery and distribution.

We have successfully integrated previously established Intracranial window (ICW) technology 1-4 with intravital 2-photon confocal microscopy to develop a ...

A Human Ex Vivo Atherosclerotic Plaque Model to Study Lesion Biology

Atherosclerosis is a chronic inflammatory disease of the vasculature. There are various methods to study the inflammatory compound in atherosclerotic lesions. Mouse models are an important tool to investigate inflammatory processes in atherogenesis, but these models suffer from the phenotypic and functional differences between the murine and human immune system. In vitro cell experiments are used to specifically evaluate cell type-dependent changes caused by a substance of interest, but culture-dependent variations and the inability to analyze the influence of specific molecules in the context of the inflammatory compound in atherosclerotic lesions limit the impact of the results. In addition, measuring levels of a molecule of interest in human blood helps to further investigate its clinical relevance, but this represents systemic and not local inflammation. Therefore, we here describe a plaque culture model to study human atherosclerotic lesion biology ex vivo. In short, fresh plaques are obtained from patients undergoing endarterectomy or coronary artery bypass grafting and stored in RPMI medium on ice until usage. The specimens are cut into small pieces followed by random distribution into a 48-well plate, containing RPMI medium in addition to a substance of interest such as cytokines or chemokines alone or in combination for defined periods of time. After incubation, the plaque pieces can be shock frozen for mRNA isolation, embedded in Paraffin or OCT for immunohistochemistry staining or smashed and lysed for western blotting. Furthermore, cells may be isolated from the plaque for flow cytometry analysis. In addition, supernatants can be collected for protein measurement by ELISA. In conclusion, the presented ex vivo model opens the possibility to further study inflammatory lesional biology, which may result in identification of novel disease mechanisms and therapeutic targets.

A Human Ex Vivo Atherosclerotic Plaque Model to Study Lesion Biology

Atherosclerosis is a chronic inflammatory process. This manuscript illustrates an easy to use ex vivo model to investigate fresh carotid or coronary artery plaques. The ex vivo model allows for the investigation of potential substances on the inflammatory milieu in human atherosclerotic lesions and results can be analyzed by various methods.

Atherosclerosis is a chronic inflammatory disease of the vasculature. There are various methods to study the inflammatory compound in atherosclerotic ...

In vivo 19F MRI for Cell Tracking

In vivo 19F MRI allows quantitative cell tracking without the use of ionizing radiation. It is a noninvasive technique that can be applied to humans. Here, we describe a general protocol for cell labeling, imaging, and image processing. The technique is applicable to various cell types and animal models, although here we focus on a typical mouse model for tracking murine immune cells. The most important issues for cell labeling are described, as these are relevant to all models. Similarly, key imaging parameters are listed, although the details will vary depending on the MRI system and the individual setup. Finally, we include an image processing protocol for quantification. Variations for this, and other parts of the protocol, are assessed in the Discussion section. Based on the detailed procedure described here, the user will need to adapt the protocol for each specific cell type, cell label, animal model, and imaging setup. Note that the protocol can also be adapted for human use, as long as clinical restrictions are met.

In vivo 19F MRI for Cell Tracking

We describe a general protocol for in vivo cell tracking using MRI in a mouse model with ex vivo labeled cells. A typical protocol for cell labeling, image acquisition processing and quantification is included.

In vivo 19F MRI allows quantitative cell tracking without the use of ionizing radiation. It is a noninvasive technique that can be applied to humans. ...

Molecular Profiling of the Invasive Tumor Microenvironment in a 3-Dimensional Model of Colorectal Cancer Cells and Ex vivo Fibroblasts

Invading colorectal cancer (CRC) cells have acquired the capacity to break free from their sister cells, infiltrate the stroma, and remodel the extracellular matrix (ECM). Characterizing the biology of this phenotypically distinct group of cells could substantially improve our understanding of early events during the metastatic cascade.

Tumor invasion is a dynamic process facilitated by bidirectional interactions between malignant epithelium and the cancer associated stroma. In order to examine cell-specific responses at the tumor stroma-interface we have combined organotypic co-culture and laser micro-dissection techniques.
Organotypic models, in which key stromal constituents such as fibroblasts are 3-dimentioanally co-cultured with cancer epithelial cells, are highly manipulatable experimental tools which enable invasion and cancer-stroma interactions to be studied in near-physiological conditions.

Laser microdissection (LMD) is a technique which entails the surgical dissection and extraction of the various strata within tumor tissue, with micron level precision.
By combining these techniques with genomic, transcriptomic and epigenetic profiling we aim to develop a deeper understanding of the molecular characteristics of invading tumor cells and surrounding stromal tissue, and in doing so potentially reveal novel biomarkers and opportunities for drug development in CRC.&#160;&#160;&#160;

Molecular Profiling of the Invasive Tumor Microenvironment in a 3-Dimensional Model of Colorectal Cancer Cells and Ex vivo Fibroblasts

Molecular profiling of laser microdissected cells and stroma from synthetic, tissue-engineered colorectal cancer models represents a novel and manipulatable approach to characterize and dissect the distinctive biology at the interface between tumor cells at the invasive front and cancer associated stromal cells. &#160;

Invading colorectal cancer (CRC) cells have acquired the capacity to break free from their sister cells, infiltrate the stroma, and remodel the ...

An Ex vivo Model to Study Hormone Action in the Human Breast

The study of hormone action in the human breast has been hampered by lack of adequate model systems. Upon in vitro culture, primary mammary epithelial cells tend to lose hormone receptor expression. Widely used hormone receptor positive breast cancer cell lines are of limited relevance to the in vivo situation. Here, we describe an ex vivo model to study hormone action in the human breast. Fresh human breast tissue specimens from surgical discard material such as reduction mammoplasties or mammectomies are mechanically and enzymatically digested to obtain tissue fragments containing ducts and lobules and multiple stromal cell types. These tissue microstructures kept in basal medium without growth factors preserve their intercellular contacts, the tissue architecture, and remain hormone responsive for several days. They are readily processed for RNA and protein extraction, histological analysis or stored in freezing medium. Fluorescence activated cell sorting (FACS) can be used to enrich for specific cell populations. This protocol provides a straightforward, standard approach for translational studies with highly complex, varied human specimens.

An Ex vivo Model to Study Hormone Action in the Human Breast

We have developed a novel ex vivo model to study hormone action in the human breast. It is based on tissue microstructures isolated from surgical breast tissue specimens which preserve tissue architecture, intercellular interactions, and paracrine signaling.

The study of hormone action in the human breast has been hampered by lack of adequate model systems. Upon in vitro culture, primary mammary epithelial ...

Validation of Nanobody and Antibody Based In Vivo Tumor Xenograft NIRF-imaging Experiments in Mice Using Ex Vivo Flow Cytometry and Microscopy

This protocol outlines the steps required to perform ex vivo validation of in vivo near-infrared fluorescence (NIRF) xenograft imaging experiments in mice using fluorophore labelled nanobodies and conventional antibodies.
First we describe how to generate subcutaneous tumors in mice, using antigen-negative cell lines as negative controls and antigen-positive cells as positive controls in the same mice for intraindividual comparison. We outline how to administer intravenously near-infrared fluorophore labelled (AlexaFluor680) antigen-specific nanobodies and conventional antibodies. In vivo imaging was performed with a small-animal NIRF-Imaging system. After the in vivo imaging experiments the mice were sacrificed. We then describe how to prepare the tumors for parallel ex vivo analyses by flow cytometry and fluorescence microscopy to validate in vivo imaging results.
The use of the near-infrared fluorophore labelled nanobodies allows for non-invasive same day imaging in vivo. Our protocols describe the ex vivo quantification of the specific labeling efficiency of tumor cells by flow cytometry and analysis of the distribution of the antibody constructs within the tumors by fluorescence microscopy. Using near-infrared fluorophore labelled probes allows for non-invasive, economical in vivo imaging with the unique ability to exploit the same probe without further secondary labelling for ex vivo validation experiments using flow cytometry and fluorescence microscopy.

Validation of Nanobody and Antibody Based In Vivo Tumor Xenograft NIRF-imaging Experiments in Mice Using Ex Vivo Flow Cytometry and Microscopy

This protocol outlines the steps required to perform ex vivo validation of in vivo near-infrared fluorescence xenograft imaging experiments in mice using fluorophore labelled nanobodies and conventional antibodies.

This protocol outlines the steps required to perform ex vivo validation of in vivo near-infrared fluorescence (NIRF) xenograft imaging experiments in mice ...

Human Dupuytren's Ex Vivo Culture for the Study of Myofibroblasts and Extracellular Matrix Interactions

Organ fibrosis or &#8220;scarring&#8221; is known to account for a high death toll due to the extensive amount of disorders and organs affected (from cirrhosis to cardiovascular diseases). There is no effective treatment and the in vitro tools available do not mimic the in vivo situation rendering the progress of the out of control wound healing process still enigmatic.
To date, 2D and 3D cultures of fibroblasts derived from DD patients are the main experimental models available. Primary cell cultures have many limitations; the fibroblasts derived from DD are altered by the culture conditions, lack cellular context and interactions, which are crucial for the development of fibrosis and weakly represent the derived tissue. Real-time PCR analysis of fibroblasts derived from control and DD samples show that little difference is detectable. 3D cultures of fibroblasts include addition of extracellular matrix that alters the native conditions of these cells. 
As a way to characterize the fibrotic, proliferative properties of these resection specimens we have developed a 3D culture system, using intact human resections of the nodule part of the cord. The system is based on transwell plates with an attached nitrocellulose membrane that allows contact of the tissue with the medium but not with the plastic, thus, preventing the alteration of the tissue. No collagen gel or other extracellular matrix protein substrate is required. The tissue resection specimens maintain their viability and proliferative properties for 7 days. This is the first &#8220;organ&#8221; culture system that allows human resection specimens from DD patients to be grown ex vivo and functionally tested, recapitulating the in vivo situation.

Human Dupuytren&#039;s Ex Vivo Culture for the Study of Myofibroblasts and Extracellular Matrix Interactions

Dupuytren&#8217;s disease (DD) is a fibroproliferative disease of the palm of the hand. Here, we present a protocol to culture resection specimens from DD in a three-dimensional (3D) culture system. Such short-term culture system allows preservation of the 3D structure and molecular properties of the fibrotic tissue.

Organ fibrosis or &#8220;scarring&#8221; is known to account for a high death toll due to the extensive amount of disorders and organs affected (from ...

Intrauterine Telemetry to Measure Mouse Contractile Pressure In Vivo

A complex integration of molecular and electrical signals is needed to transform a quiescent uterus into a contractile organ at the end of pregnancy. Despite the discovery of key regulators of uterine contractility, this process is still not fully understood. Transgenic mice provide an ideal model in which to study parturition. Previously, the only method to study uterine contractility in the mouse was ex vivo isometric tension recordings, which are suboptimal for several reasons. The uterus must be removed from its physiological environment, a limited time course of investigation is possible, and the mice must be sacrificed. The recent development of radiometric telemetry has allowed for longitudinal, real-time measurements of in vivo intrauterine pressure in mice. Here, the implantation of an intrauterine telemeter to measure pressure changes in the mouse uterus from mid-pregnancy until delivery is described. By comparing differences in pressures between wild type and transgenic mice, the physiological impact of a gene of interest can be elucidated. This technique should expedite the development of therapeutics used to treat myometrial disorders during pregnancy, including preterm labor.

Intrauterine Telemetry to Measure Mouse Contractile Pressure In Vivo

This manuscript provides a protocol for implanting telemeters in the mouse for the purpose of measuring intrauterine pressures during pregnancy.

A complex integration of molecular and electrical signals is needed to transform a quiescent uterus into a contractile organ at the end of pregnancy. ...