The authors thank Dr. Laura L. Bronsart for providing the GFP- and luciferase-4T1 cells, Dr. Edward L. LaGory for advice on 1-([4-(Xylylazo)xylyl]azo)-2-naphthol staining, Dr. Craig L. Duvall for IVIS and lyophilizer use, and Dr. Scott A. Guelcher for rheometer use. This research was financially supported by NIH grant #R00CA201304.

Animal studies were performed in accordance with institutional guidelines and protocols approved by the Vanderbilt University Institutional Animal Care and Use Committee.
1. Preparation and ex vivo irradiation of MFPs
<ol>
	<li>Sacrifice athymic Nu/Nu mice (8&#8211;10 weeks) using CO&#173;2 asphyxiation followed by cervical dislocation.</li>
	<li>Clean the skin using 70% ethanol.</li>
	<li>Collect mammary fat pads (MFPs) from sacrificed mice using pre-sterilized scissors and forceps in a 15 mL conical tube containing complete RPMI media (RPMI supplemented with 1% penicillin-streptomycin and 10% fetal bovine serum) (see the Table of Materials).</li>
	<li>Irradiate samples to 20 Gy using a cesium source.</li>
	<li>Bring the irradiated MFPs and complete RPMI media into a biosafety cabinet. The media will be dependent on the cell line to be grown in the final hydrogel. Fill 6 cm or 10 cm dishes with enough media to submerge the MFPs. For 6 cm dishes, use 8 mL of media, and for 10 cm dishes, use 20 mL of media.</li>
	<li>Incubate in a 37 &#176;C/5% CO2 incubator for two days. The length of time in the incubator may be adjusted.</li>
	<li>Rinse tissues in phosphate-buffered saline (PBS), blot excess moisture, and place MFPs into 15 mL conical tubes for storage at -80 &#176;C until decellularization. This freezing step aids the decellularization step and the sample should be frozen even if the next steps are otherwise ready.</li>
</ol>
2. Decellularization (adapted from references12,16,17)
NOTE: This procedure was adapted from previously published methods focused on adipose decellularization, which included the sodium deoxycholate ionic detergent rather than sodium dodecyl sulfate to remove DNA efficiently12,16,17.
<ol>
	<li>On day 1, remove frozen MFPs from -80 &#176;C and thaw at room temperature.</li>
	<li>Once thawed, dry MFPs briefly on a delicate task wipe. Weigh the MFPs using an analytical scale.</li>
	<li>Using a pair of forceps with scissors or a scalpel, divide tissue up into 3 mm x 3 mm x 3 mm samples for study of the intact ECM and the remaining tissue for hydrogel production. 
	NOTE: The number of samples is dependent on the number of testing methods, e.g. the collection of two samples is described below: one for paraffin embedding (step 2.5) and one for freezing in cryostat embedding medium, if desired (see the Table of Materials and step 2.6).</li>
	<li>Weigh the tissues. If embedding in paraffin for sectioning, continue to step 2.5. If freezing in cryostat embedding medium for sectioning, continue to step 2.6.</li>
	<li>In a chemical hood, submerge the tissue in 10% neutral buffered formalin (NBF) (see the Table of Materials) for 24 h at 4 &#176;C. Wash 3 times in PBS for 5 min each. Submerge the tissue in 30% sucrose for 48 h at 4 &#176;C.
	<ol>
		<li>Weigh the tissue now that this piece has been removed. Continue to step 2.6.</li>
	</ol>
	</li>
	<li>In a chemical hood, place MFP pieces in a labeled cassette prepped with cryostat embedding medium. Add more cryostat embedding medium to cover the tissue.
	<ol>
		<li>Place the cassette into a beaker of 2-methylbutane (see the Table of Materials) that is pre-cooled with liquid nitrogen. The beaker should have enough 2-methylbutane to cover the bottom but not enough to submerge the cassette because the cryostat embedding medium should not touch the 2-methylbutane. Let the cassette sit in the 2-methylbutane until the cryostat embedding medium freezes and becomes opaque.</li>
		<li>Wrap the cassette(s) in foil, label, and leave at -80 &#176;C until used for sectioning. 
		NOTE: Tissues placed immediately in cryostat embedding medium were sectioned at 5 &#956;m while tissues incubated in sucrose were sectioned at 30 &#956;m to retain adipocyte morphology.</li>
	</ol>
	</li>
	<li>Use forceps to manually massage the remaining tissue. 
	NOTE: Tissue pieces may also be placed in 10% NBF for 24&#8211;48 h, rinsed in PBS, and left in 70% ethanol until embedding in paraffin. Following embedding, 5 &#956;m sections can be used for hematoxylin and eosin (H &#38; E) staining (see section 7 below).</li>
	<li>Place the MFPs in 6 cm dishes with 5 mL 0.02% trypsin/0.05% EDTA solution. Incubate at 37 &#176;C for 1 h. Spray and wipe the dishes with 70% ethanol before placing in the incubator.</li>
	<li>Use 0.7 mm strainers to wash the MFPs with deionized (DI) water by pouring water over the tissue three times. Use forceps to manually massage the tissue in between washes.</li>
	<li>Briefly dry tissue on a delicate task wipe and weigh. Place tissues in a pre-autoclaved beaker containing an appropriately sized stir bar. Cover tissues with 60 mL of 3% t-octylphenoxypolyethoxyethanol (see the Table of Materials) per 1 g of tissue and stir for 1 h at room temperature. Use a minimum of 20 mL.</li>
	<li>Dump tissue and contents into a strainer. Rinse the beaker with&#160;DI&#160;water and pour onto tissues. Repeat two more times. Use forceps to manually massage the tissue in between rinses.</li>
	<li>Briefly dry tissue on a delicate task wipe and weigh. Place tissues and stir bars back in the same beakers, and cover with 60 mL of 4% deoxycholic acid per 1 g of tissue. Stir for 1 h at room temperature. Use a minimum of 20 mL.</li>
	<li>Dump tissue and contents into a mesh strainer. Rinse the beaker with DI water and pour onto tissues. Repeat two more times. Use forceps to manually massage the tissue in between rinses.</li>
	<li>Briefly dry tissue on a delicate task wipe and weigh.</li>
	<li>Place tissues in the same beaker with fresh DI water supplemented with 1% penicillin-streptomycin. Cover tightly with paraffin film. Leave overnight at 4 &#176;C.</li>
	<li>Wash strainers and beakers for use the following day.</li>
	<li>On day 2, drain beaker contents into a strainer. Briefly dry tissue on a delicate task wipe and weigh.</li>
	<li>Place MFPs in the same beaker with an appropriately sized stir bar. Cover with 60 mL 4% ethanol/0.1% peracetic acid solution per 1 g of tissue. Use a minimum of 20 mL. Stir for 2 h at room temperature.</li>
	<li>Dump tissue and contents into a 0.7 mm strainer. Use forceps to manually massage the tissue. Place contents back into the beaker. Wash tissue by covering it with 60 mL of 1x PBS per 1 g of tissue. Use a minimum of 20 mL. Stir for 15 min at room temperature. Repeat once.</li>
	<li>Dump tissue and contents into a 0.7 mm strainer. Use forceps to manually massage the tissue. Place contents back into beaker. Wash tissue by covering it with 60 mL DI water per 1 g of tissue. Use a minimum of 20 mL. Stir for 15 min at room temperature. Repeat once.</li>
	<li>Briefly dry tissue on a delicate task wipe and weigh. Dump tissue and contents into a strainer. Use forceps to manually massage the tissue.</li>
	<li>Place contents back into beaker. Cover tissues with 60 mL of 100% n-propanol per 1 g of tissue. Use a minimum of 20 mL. Stir for 1 h at room temperature.</li>
	<li>Briefly dry tissue on a delicate task wipe and weigh. Dump tissue and contents into a 0.7 mm strainer. Use forceps to manually massage the tissue.</li>
	<li>Place contents back into beaker. Wash tissue by covering it with 60 mL of DI water per 1 g of tissue. Use a minimum of 20 mL. Stir for 15 min at room temperature. Repeat three times.</li>
	<li>Dump tissue and contents into a strainer. Repeat steps 2.3&#8211;2.6 to collect pieces of tissue for sucrose incubation and freezing in cryostat embedding medium.</li>
	<li>Briefly dry tissue on a delicate task wipe and weigh. Place in a labeled 15 mL tube. Freeze at -80 &#176;C overnight.</li>
</ol>
3. Lyophilization
<ol>
	<li>Remove the 15 mL tubes from -80 &#176;C and place on dry ice. Keep samples frozen on dry ice until on the lyophilizer.</li>
	<li>Remove caps. Use a rubber band to attach a delicate task wipe to the top to cover the opening. Put samples on the lyophilizer for at least 2 days.</li>
	<li>Remove samples from the lyophilizer and place tubes on dry ice. Remove delicate task wipes weigh each sample on an analytical scale. Attach caps and place at -80 &#176;C overnight.</li>
</ol>
4. Milling
<ol>
	<li>Fill a shallow container with liquid nitrogen. Remove samples from the -80 &#176;C freezer. Weigh each lyophilized MFP.</li>
	<li>Place one sample in the mortar. Use a cryogenic glove to hold the mortar in the liquid nitrogen.</li>
	<li>Use a pestle attached to a handheld drill to mill the sample. Mill in 1 min intervals to check progress and remove gloved hand from liquid nitrogen. Mill for a minimum of 5 min.</li>
	<li>Repeat for all samples. Spray and wipe the mortar and pestle with ethanol between each sample.</li>
	<li>Store powdered samples in 15 mL tubes at -80 &#176;C until ready for use. 
	NOTE: Samples may be used immediately or stored overnight.</li>
</ol>
5. Hydrogel formation
<ol>
	<li>If stored at -80 &#176;C overnight, remove and thaw at room temperature. While thawing, calculate the necessary weight of pepsin (see Table of Materials) and volume of hydrochloric acid (HCl) needed for each sample that results in a solution with 1% w/v sample powder and 0.1% w/v pepsin in 0.01 M HCl. Add the pepsin to the HCl to form a pepsin-HCl solution.</li>
	<li>Add sample powder and pepsin-HCl solution to a 15 mL tube. Add a small stir bar, and stir for 48 h.</li>
	<li>Place the tubes on ice for 5 min. Calculate the necessary volume of 10x PBS needed for each sample that results in a solution with a 1x PBS concentration. Add the appropriate volume of 10x PBS to each tube.</li>
	<li>Add 10% v/v 0.1 M NaOH to each solution to reach pH 7.4. Use a pH paper&#160;to test individually. 
	NOTE: Gel solution may be stored at 4 &#176;C for one week.</li>
</ol>
6. Encapsulating cells in hydrogels
<ol>
	<li>Using GFP- and luciferase-labeled cells
	<ol>
		<li>Using the pH 7.4 gel solution, resuspend pelleted GFP- and luciferase-labeled 4T1 cells to a concentration of 500,000 or 1,000,000 cells/mL of gel solution. Add 16 &#181;L of gel-cell solution to each well of a 16-well chamber slide. Incubate for 30 min at 37 &#176;C.</li>
		<li>Add 100 &#181;L of complete RPMI media to each well. Continue incubation for 48 h at 37 &#176;C. The GFP- and luciferase-labeled cells can be visualized using fluorescence microscopy at 0 hours, 24 hours, and 48 hours after gelation. 
		NOTE: Cell proliferation can be measured for the GFP- and luciferase-labeled 4T1 cells used here by adding (S)-4,5-Dihydro-2-(6-hydroxy-2-benzothiazolyl)-4-thiazolecarboxylic acid potassium salt (see the Table of Materials) to the wells for 10 min and performing bioluminescence imaging using a bioluminescence imaging system (see Table of Materials). Following bioluminescence imaging, the cytoskeleton can be visualized (see section 10).</li>
	</ol>
	</li>
	<li>Using unlabeled cells
	<ol>
		<li>Using the pH 7.4 gel solution, resuspend pelleted unlabeled 4T1 cells to a concentration of 500,000 or 1,000,000 cells/mL of gel solution. Add 16 &#181;L of gel-cell solution to each well of a 16-well chamber slide and a 96-well plate. Incubate for 30 min at 37 &#176;C.</li>
		<li>Add 100 &#181;L of media to each well. Continue incubation for 48 h at 37 &#176;C. 
		NOTE: Cell viability can be measured (see section 11). The 16-well chamber slide can be used for imaging, and the 96-well plate can be used for quantification.</li>
	</ol>
	</li>
</ol>
7. H &#38; E staining
<ol>
	<li>For formalin-fixed, paraffin-embedded tissue, submerge slides containing 5 &#956;m sections twice in 100% xylene (5&#8211;10 min) for de-paraffinization. Rehydrate by submerging in 100%, 95%, 85%, and 70% ethanol for 5 min each followed by DI water for 5 min. Continue to step 7.6.</li>
	<li>For frozen tissue sections, take sections immediately from freezer and submerge them in 10% NBF for 10 min. Wash in 1x PBS three times for 5 min each. Rinse in water for 5 min.</li>
	<li>Stain nuclei with hematoxylin for 3 min. Rinse in running tap water.</li>
	<li>Differentiate by dipping 1&#8211;2 times in 0.3% acid alcohol (0.3% HCl in 70% ethanol). Rinse in running tap water for 5 min.</li>
	<li>Add bluing agent for 30 s. Rinse in running tap water for 5 min. Submerge in 95% ethanol for 30 s.</li>
	<li>Incubate with eosin for 90 s at room temperature. Dehydrate in 3 changes of 100% ethanol for 5 min each.</li>
	<li>Submerge twice in 100% xylene for 5 min each. Add distyrene-plasticiser-xylene (DPX0 mounting media to the slide and add a coverslip. Let it cure overnight before imaging.</li>
</ol>
8. 1-([4-(Xylylazo)xylyl]azo)-2-naphthol staining
<ol>
	<li>Prepare 1-([4-(Xylylazo)xylyl]azo)-2-naphthol solution.
	<ol>
		<li>Add 0.5 g of 1-([4-(Xylylazo)xylyl]azo)-2-naphthol powder to a beaker with 100 mL propylene glycol. Heat to 95&#8211;100 &#176;C while stirring for at least 30 min. Prevent the temperature from exceeding 100 &#176;C.</li>
		<li>Remove beaker from heat and allow to cool slightly. Pour solution through grade 4 qualitative filter paper to remove any residual particulates. 
		NOTE: The solution can be stored at room temperature and should be filtered through a 0.45 &#181;m syringe filter immediately before staining.</li>
	</ol>
	</li>
	<li>Stain frozen tissue sections.
	<ol>
		<li>Remove slides from the -80 &#176;C freezer and air dry for 30 min. Pre-cool 10% NBF at -20 &#176;C for 30 min in a Coplin jar. Fix the slides at room temperature for 10 min. Wash in DI water 3 times for 5 min each.</li>
		<li>Remove excess water using a delicate task wipe before immersing in propylene glycol 2 times for 5 min each. Remove slides from propylene glycol. Do not rinse.</li>
		<li>Place slides into syringe filtered Oil Red O staining solution for 3 h at room temperature.</li>
		<li>Differentiate by placing slides in 85% propylene glycol for 5 min. Rinse samples in PBS twice for 5 min each.</li>
		<li>Stain samples with hematoxylin for 30 s. Wash in running tap water for 5 min and then place the slides in DI water.</li>
		<li>Mount samples using aqueous based mounting medium and allow media to dry overnight before imaging.</li>
	</ol>
	</li>
</ol>
9. Rheology
<ol>
	<li>Bring pre-gel solutions from step 5.4 to the rheometer on ice.</li>
	<li>Attach a 25 mm arm and plate to the rheometer. Open the rheology software on the computer connected to the rheometer.</li>
	<li>Perform rotational mapping and determine the zero gap. Raise the arm when complete.</li>
	<li>Pipette 500 &#956;L of pre-gel on the plate. Lower the arm until the pre-gel solution completely occupies the gap between the plate and the arm. Be conservative because lowering the arm past this point will result in pre-gel solution spilling out of the side.</li>
	<li>Perform a frequency sweep on the pre-gel solution from 0.1&#8211;100 Hz after it has remained at 37 &#176;C for 30 min.</li>
	<li>Raise the rheometer arm when complete. Wipe liquid with a delicate task wipe. Rinse with DI water and wipe with a delicate task wipe. Repeat steps 9.4&#8211;9.6 with additional samples.</li>
</ol>
10. Phalloidin conjugate staining of F-actin
<ol>
	<li>Bring phalloidin conjugate solution to room temperature. Briefly centrifuge at a low speed prior to opening. Aliquot enough solution for the assay, and store the rest at -20 &#176;C.</li>
	<li>Dilute 1000x phalloidin conjugate stock solution to a 1x working solution by adding 1 &#956;L stock solution to 1 mL 1x PBS + 1% bovine serum albumin (BSA). This makes enough for 10 wells (100 &#956;L/well). 
	NOTE: One may use plain 1x PBS, but 1x PBS + BSA is preferred to prevent phalloidin conjugate from sticking to the tube. Do not store this solution after the assay. 1 &#956;L of blue fluorescent dye (see the Table of Materials) working solution may be added to stain for nuclei. Working solution may be made by mixing 1 &#956;L of 20 mM blue fluorescent dye with 11.3 &#956;L 1x PBS.</li>
	<li>Aspirate media from wells (step 6.1). Rinse wells with 1x PBS.</li>
	<li>Add 10% NBF. Incubate wells with 10% NBF for 20 min at room temperature. Remove supernatant. Wash 3 times with PBS for 5 min each time.</li>
	<li>Add 0.1% t-Octylphenoxypolyethoxyethanol in PBS for 5 min. Aspirate and wash 3 times with PBS for 5 min each time.</li>
	<li>Add 100 &#956;L of working solution from step 10.2 to each well. Incubate for 1 h at room temperature. Aspirate and wash 3 times with PBS for 5 min each time. Leave in PBS at 4 &#176;C.</li>
	<li>Aspirate and remove wells from the gasket on the chamber slide. Use needle nose tweezers to remove the gasket from the slide. Mount and coverslip samples using aqueous based mounting medium. Allow to cure (5 min).</li>
	<li>Observe cells under a fluorescence microscope at excitation/emission of 590/618 nm.</li>
</ol>
11. Viability assay
<ol>
	<li>Rinse the cells with Dulbecco&#8217;s PBS (DPBS). Add 100 &#181;L of DPBS containing 1 &#181;M calcein AM and 2 &#181;M ethidium homodimer to each well. Incubate for 30 min at room temperature.</li>
	<li>Image the 16-well chamber slide by fluorescence microscopy. Calcein acetoxymethyl (calcein AM) can be observed at excitation/emission = 494/517 nm. Ethidium homodimer can be observed at excitation/emission = 528/645 nm.</li>
	<li>Read the 96-well plate on a plate reader using the same wavelengths in step 11.2.</li>
</ol>

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T., Frey, B., Fellner, M., Herrmann, M., Fabry, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Integrin+5+1+facilitates+cancer+cell+invasion+through+enhanced+contractile+forces.">Integrin 5 1 facilitates cancer cell invasion through enhanced contractile forces.</a> Journal of Cell Science. 124 (3), 369-383 (2011).</li><li>Ahmadzadeh, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modeling+the+two-way+feedback+between+contractility+and+matrix+realignment+reveals+a+nonlinear+mode+of+cancer+cell+invasion.">Modeling the two-way feedback between contractility and matrix realignment reveals a nonlinear mode of cancer cell invasion.</a> Proceedings of the National Academy of Sciences. 114 (9), E1617-E1626 (2017).</li></ol>

The authors have nothing to disclose.

This method of hydrogel formation is largely dependent on the amount of starting tissue. Murine MFPs are small, and the decellularization process results in a significant reduction of material (Table 1). The process can be repeated with more MFPs to increase final yield. Milling is another important step that may lead to loss of material. Others have shown success with a cryogenic mill, but this protocol is based on milling via a handheld mortar and electric drill with a pestle attachment<sup class="xref...

Cancer is characterized by excess proliferation of cells that can evade apoptosis and also metastasize to distant sites1. Breast cancer is one of the most common forms among females in the US, with an estimated 266,000 new cases and 40,000 deaths in 20182. A particularly aggressive and difficult to treat subtype is triple negative breast cancer (TNBC), which lacks estrogen receptor (ER), progesterone receptor (PR), and human epidermal growth factor (HER2). Radiation therapy is commonly used in breast cancer to eliminate residual tumor cells following lumpectomy, but over 13% of TNBC patients still experience recurrence at the primary tumor site3.
It is known that radiation therapy is effective in mitigating metastasis and recurrence because the combination of lumpectomy and radiation results in the same long-term survival as mastectomy4. However, it has recently been shown that radiation treatment is associated with local recurrence to the primary tumor site in immunocompromised settings5,6. Also, it is well known that radiation changes the extracellular matrix (ECM) of normal tissue by inducing fibrosis7. Therefore, it is important to understand the role of radiation-induced ECM changes in dictating tumor cell behavior.
Decellularized tissues have been used as in vitro models to study disease8,9. These decellularized tissues preserve ECM composition and recapitulate the complex in vivo ECM. This decellularized tissue ECM can be further processed and digested to form reconstituted ECM hydrogels that can be used to study cell growth and function10,11. For example, injectable hydrogels derived from decellularized human lipoaspirate and from myocardial tissue served as non-invasive methods of tissue engineering, and a hydrogel derived from porcine lung tissue was utilized as an in vitro method of testing mesenchymal stem cell attachment and viability12,13,14. The effect of normal tissue radiation damage on ECM properties, however, has not been investigated.
Hydrogels derived from ECM have the greatest potential for in vitro study of in vivo phenomena. Several other materials have been studied, including collagen, fibrin, and matrigel, but it is difficult to synthetically recapitulate the composition of the ECM13. An advantage of using ECM-derived hydrogels is that the ECM contains the necessary proteins and growth factors for a particular tissue14,15. Irradiation of normal tissue during lumpectomy causes significant changes to the ECM, and ECM-derived hydrogels can be used to study this effect in vitro. This method could lead to more complex and more accurate in vitro models of disease.
In this study, we subjected murine mammary fat pads (MFPs) to radiation ex vivo. The MFPs were decellularized and made into pre-gel solution. Hydrogels were formed with embedded 4T1 cells, a murine TNBC cell line. The rheological properties of the hydrogel material were examined, and tumor cell dynamics were evaluated within the hydrogels. Hydrogels fabricated from irradiated MFPs enhanced tumor cell proliferation. Future studies will incorporate other cell types to study cell-cell interactions in the context of cancer recurrence following therapy.

<table>
	<tbody>
		<tr>
			<td>10% Neutral Buffered Formalin, Cube with Spigot</td>
			<td>VWR</td>
			<td>16004-128</td>
			<td>-</td>
		</tr>
		<tr>
			<td>2-methylbutane</td>
			<td>Alfa Aesar</td>
			<td>19387</td>
			<td>-</td>
		</tr>
		<tr>
			<td>AR 2000ex Rheometer</td>
			<td>TA Instruments</td>
			<td>10D4335</td>
			<td>rheometer</td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Sigma-Aldrich</td>
			<td>A1933-25G</td>
			<td>-</td>
		</tr>
		<tr>
			<td>calcein acetoxymethyl (calcein AM)</td>
			<td>Molecular Probes, Inc.</td>
			<td>C1430</td>
			<td>-</td>
		</tr>
		<tr>
			<td>D-Luciferin Firefly, potassium salt</td>
			<td>Biosynth Chemistry &#38; Biology</td>
			<td>L-8820</td>
			<td>(S)-4,5-Dihydro-2-(6-hydroxy-2-benzothiazolyl)-4-thiazolecarboxylic acid potassium salt</td>
		</tr>
		<tr>
			<td>DPX Mountant for Histology</td>
			<td>Sigma-Aldrich</td>
			<td>06522-500ML</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s phosphate-buffered saline</td>
			<td>Gibco</td>
			<td>14040133</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Eosin-Y with Phloxine</td>
			<td>Richard-Allan Scientific</td>
			<td>71304</td>
			<td>eosin</td>
		</tr>
		<tr>
			<td>ethidium homodimer</td>
			<td>Molecular Probes, Inc.</td>
			<td>E1169</td>
			<td>ethidium homodimer-1 (EthD-1)</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Sigma-Aldrich</td>
			<td>F0926-500ML</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Fisher Healthcare Tissue-Plus O.C.T. Compound</td>
			<td>Fisher Scientific</td>
			<td>23-730-571</td>
			<td>cryostat embedding medium</td>
		</tr>
		<tr>
			<td>Fluoromount-G</td>
			<td>SouthernBiotech</td>
			<td>0100-01</td>
			<td>aqueous based mounting medium</td>
		</tr>
		<tr>
			<td>FreeZone 4.5</td>
			<td>Labconco</td>
			<td>7751020</td>
			<td>lyophilizer</td>
		</tr>
		<tr>
			<td>Hoechst 33342 Solution (20 mM)</td>
			<td>Thermo Scientific</td>
			<td>62249</td>
			<td>blue fluorescent dye</td>
		</tr>
		<tr>
			<td>Hydrochloric acid</td>
			<td>Sigma-Aldrich</td>
			<td>258148-500ML</td>
			<td>-</td>
		</tr>
		<tr>
			<td>IVIS Lumina III</td>
			<td>PerkinElmer</td>
			<td>CLS136334</td>
			<td>bioluminescence imaging system</td>
		</tr>
		<tr>
			<td>Kimtech Science Kimwipes</td>
			<td>Kimberly Clark</td>
			<td></td>
			<td>delicate task wipes</td>
		</tr>
		<tr>
			<td>n-Propanol (Peroxide-Free/Sequencing), Fisher BioReagents</td>
			<td>Fisher Scientific</td>
			<td>BP1130-500</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Oil Red O</td>
			<td>Sigma-Aldrich</td>
			<td>O0625-25G</td>
			<td>1-([4-(Xylylazo)xylyl]azo)-2-naphthol</td>
		</tr>
		<tr>
			<td>OPS Diagnostics CryoGrinder</td>
			<td>OPS Diagnostics, LLC</td>
			<td>CG-08-02</td>
			<td>-</td>
		</tr>
		<tr>
			<td>PBS (10X), pH 7.4</td>
			<td>Quality Biological, Inc.</td>
			<td>119-069-151</td>
			<td>Phosphate-buffered saline</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Gibco</td>
			<td>15140-122</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Pepsin from porcine gastric mucosa</td>
			<td>Sigma-Aldrich</td>
			<td>P6887-5G</td>
			<td>pepsin</td>
		</tr>
		<tr>
			<td>Peracetic acid</td>
			<td>Sigma-Aldrich</td>
			<td>77240-100ML</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Phalloidin-iFluor 594 Reagent (ab176757)</td>
			<td>abcam</td>
			<td>ab176757</td>
			<td>phalloidin conjugate</td>
		</tr>
		<tr>
			<td>Propylene glycol</td>
			<td>Sigma-Aldrich</td>
			<td>W294004-1KG-K</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Richard-Allan Scientific Signature Series Bluing Reagent</td>
			<td>Richard-Allan Scientific</td>
			<td>7301L</td>
			<td>bluing agent</td>
		</tr>
		<tr>
			<td>Richard-Allan Scientific Signature Series Hematoxylin 7211</td>
			<td>Richard-Allan Scientific</td>
			<td>7211</td>
			<td>-</td>
		</tr>
		<tr>
			<td>RPMI Medium 1640</td>
			<td>Gibco</td>
			<td>11875-093</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Sodium deoxycholate, 98%</td>
			<td>Frontier Scientific</td>
			<td>JK559522</td>
			<td>deoxycholic acid</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma-Aldrich</td>
			<td>S5016</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Triton x-100</td>
			<td>Sigma-Aldrich</td>
			<td>X100-100ML</td>
			<td>t-Octylphenoxypolyethoxyethanol</td>
		</tr>
		<tr>
			<td>Trypsin-EDTA (0.25%), phenol red</td>
			<td>Gibco</td>
			<td>25200-056</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Whatman qualitative filter paper, Grade 4</td>
			<td>Whatman</td>
			<td>1004-110</td>
			<td>grade 4 qualitative filter paper</td>
		</tr>
		<tr>
			<td>Xylenes (Certified ACS), Fisher Chemical</td>
			<td>Fisher Scientific</td>
			<td>X5-4</td>
			<td>-</td>
		</tr>
	</tbody>
</table>

studying normal tissue radiation effects using extracellular matrix hydrogels

Radiation is a therapy for patients with triple negative breast cancer. The effect of radiation on the extracellular matrix (ECM) of healthy breast tissue and its role in local recurrence at the primary tumor site are unknown. Here we present a method for the decellularization, lyophilization, and fabrication of ECM hydrogels derived from murine mammary fat pads. Results are presented on the effectiveness of the decellularization process, and rheological parameters were assessed. GFP- and luciferase-labeled breast cancer cells encapsulated in the hydrogels demonstrated an increase in proliferation in irradiated hydrogels. Finally, phalloidin conjugate staining was employed to visualize cytoskeleton organization of encapsulated tumor cells. Our goal is to present a method for fabricating hydrogels for in vitro study that mimic the in vivo breast tissue environment and its response to radiation in order to study tumor cell behavior.

MFPs were decellularized following irradiation using the procedure shown in Figure 1A. MFPs pre-decellularization (Figure 1B) and post-decellularization (Figure 1C) are shown. Decellularization was confirmed using hematoxylin and eosin (H &#38; E) staining, and 1-([4-(Xylylazo)xylyl]azo)-2-naphthol staining was used to evaluate lipid content (Figure 2). Rheological properties of the ECM hydrogels were also assessed at 37 ...

Watch this Scientific Journal Video about Studying Normal Tissue Radiation Effects using Extracellular Matrix Hydrogels at JoVE.com

Studying Normal Tissue Radiation Effects using Extracellular Matrix Hydrogels

This protocol presents a method for decellularization and subsequent hydrogel formation of murine mammary fat pads following ex vivo irradiation.

Radiation is a therapy for patients with triple negative breast cancer. The effect of radiation on the extracellular matrix (ECM) of healthy breast tissue ...

studying-normal-tissue-radiation-effects-using-extracellular-matrix

Universidad de Cartagena UDEC

Research

JoVE Journal

Cancer Research

5.8K Views.  Vanderbilt University. This protocol presents a method for decellularization and subsequent hydrogel formation of murine mammary fat pads following ex vivo irradiation.

Video: Studying Normal Tissue Radiation Effects using Extracellular Matrix Hydrogels

Analyzing the Communication Between Monocytes and Primary Breast Cancer Cells in an Extracellular Matrix Extract (ECME)-based Three-dimensional System

Embedded in the extracellular matrix (ECM), normal and neoplastic epithelial cells intimately communicate with hematopoietic and non-hematopoietic cells, thus greatly influencing normal tissue homeostasis and disease outcome. In breast cancer, tumor-associated macrophages (TAMs) play a critical role in disease progression, metastasis, and recurrence; therefore, understanding the mechanisms of monocyte chemoattraction to the tumor microenvironment and their interactions with tumor cells is important to control the disease. Here, we provide a detailed description of a three-dimensional (3D) co-culture system of human breast cancer (BrC) cells and human monocytes. BrC cells produced high basal levels of regulated on-activation, normal T-cell expressed and secreted (RANTES), monocyte chemoattractant protein-1 (MCP-1), and granulocyte-macrophage colony-stimulating factor (G-CSF), while in co-culture with monocytes, pro-inflammatory cytokines Interleukin (IL)-1 beta (IL-1&#946;) and IL-8 were enriched together with matrix metalloproteinases (MMP)-1, MMP-2, and MMP-10. This tumor stroma microenvironment promoted resistance to anoikis in MCF-10A 3D acini-like structures, chemoattraction of monocytes, and invasion of aggressive BrC cells. The protocols presented here provide an affordable alternative to study intra-tumor communication and are an example of the great potential that in vitro 3D cell systems provide to interrogate specific features of tumor biology related to tumor aggression.

Analyzing the Communication Between Monocytes and Primary Breast Cancer Cells in an Extracellular Matrix Extract ECME-based Three-dimensional System

Here, we describe a three-dimensional culture method to analyze the morphology of primary breast cancer cells, as well as to study their direct/indirect interactions with monocytes and the outcomes such as collagen degradation, immune cell recruitment, cell invasion, and promotion of cancer-related inflammation.

Embedded in the extracellular matrix (ECM), normal and neoplastic epithelial cells intimately communicate with hematopoietic and non-hematopoietic cells, ...

PET and MRI Guided Irradiation of a Glioblastoma Rat Model Using a Micro-irradiator

For decades, small animal radiation research was mostly performed using fairly crude experimental setups applying simple single-beam techniques without the ability to target a specific or well-delineated tumor volume. The delivery of radiation was achieved using fixed radiation sources or linear accelerators producing megavoltage (MV) X-rays. These devices are unable to achieve sub-millimeter precision required for small animals. Furthermore, the high doses delivered to healthy surrounding tissue hamper response assessment. To increase the translation between small animal studies and humans, our goal was to mimic the treatment of human glioblastoma in a rat model. To enable a more accurate irradiation in a preclinical setting, recently, precision image-guided small animal radiation research platforms were developed. Similar to human planning systems, treatment planning on these micro-irradiators is based on computed tomography (CT). However, low soft-tissue contrast on CT makes it very challenging to localize targets in certain tissues, such as the brain. Therefore, incorporating magnetic resonance imaging (MRI), which has excellent soft-tissue contrast compared to CT, would enable a more precise delineation of the target for irradiation. In the last decade also biological imaging techniques, such as positron emission tomography (PET) gained interest for radiation therapy treatment guidance. PET enables the visualization of e.g., glucose consumption, amino-acid transport, or hypoxia, present in the tumor. Targeting those highly proliferative or radio-resistant parts of the tumor with a higher dose could give a survival benefit. This hypothesis led to the introduction of the biological tumor volume (BTV), besides the conventional gross target volume (GTV), clinical target volume (CTV), and planned target volume (PTV).
At the preclinical imaging lab of Ghent University, a micro-irradiator, a small animal PET, and a 7 T small animal MRI are available. The goal was to incorporate MRI-guided irradiation and PET-guided sub-volume boosting in a glioblastoma rat model.

In the past, small animal irradiation was usually performed without the ability to target a well-delineated tumor volume. The goal was to mimic the treatment of human glioblastoma in rats. Using a small animal irradiation platform, we performed MRI-guided 3D conformal irradiation with PET-based sub-volume boosting in a preclinical setting.

For decades, small animal radiation research was mostly performed using fairly crude experimental setups applying simple single-beam techniques without ...

Imaging Approaches to Assessments of Toxicological Oxidative Stress Using Genetically-encoded Fluorogenic Sensors

While oxidative stress is a commonly cited toxicological mechanism, conventional methods to study it suffer from a number of shortcomings, including destruction of the sample, introduction of potential artifacts, and a lack of specificity for the reactive species involved. Thus, there is a current need in the field of toxicology for non-destructive, sensitive, and specific methods that can be used to observe and quantify intracellular redox perturbations, more commonly referred to as oxidative stress. Here, we present a method for the use of two genetically-encoded fluorogenic sensors, roGFP2 and HyPer, to be used in live-cell imaging studies to observe xenobiotic-induced oxidative responses. roGFP2 equilibrates with the glutathione redox potential (EGSH), while HyPer directly detects hydrogen peroxide (H2O2). Both sensors can be expressed into various cell types via transfection or transduction, and can be targeted to specific cellular compartments. Most importantly, live-cell microscopy using these sensors offers high spatial and temporal resolution that is not possible using conventional methods. Changes in the fluorescence intensity monitored at 510 nm serves as the readout for both genetically-encoded fluorogenic sensors when sequentially excited by 404 nm and 488 nm light. This property makes both sensors ratiometric, eliminating common microscopy artifacts and correcting for differences in sensor expression between cells. This methodology can be applied across a variety of fluorometric platforms capable of exciting and collecting emissions at the prescribed wavelengths, making it suitable for use with confocal imaging systems, conventional wide-field microscopy, and plate readers. Both genetically-encoded fluorogenic sensors have been used in a variety of cell types and toxicological studies to monitor cellular EGSH and H2O2 generation in real-time. Outlined here is a standardized method that is widely adaptable across cell types and fluorometric platforms for the application of roGFP2 and HyPer in live-cell toxicological assessments of oxidative stress.

This manuscript describes the use of genetically-encoded fluorogenic reporters in an application of live-cell imaging for the examination of xenobiotic-induced oxidative stress. This experimental approach offers unparalleled spatiotemporal resolution, sensitivity, and specificity while avoiding many of the shortcomings of conventional methods used for the detection of toxicological oxidative stress.

While oxidative stress is a commonly cited toxicological mechanism, conventional methods to study it suffer from a number of shortcomings, including ...

Fabrication of Tongue Extracellular Matrix and Reconstitution of Tongue Squamous Cell Carcinoma In Vitro

In order to construct an effective and realistic model for tongue squamous cell carcinoma (TSCC) in vitro, the methods were created to produce decellularized tongue extracellular matrix (TEM) which provides functional scaffolds for TSCC construction. TEM provides an in vitro niche for cell growth, differentiation, and cell migration. The microstructures of native extracellular matrix (ECM) and biochemical compositions retained in the decellularized matrix provide tissue-specific niches for anchoring cells. The fabrication of TEM can be realized by deoxyribonuclease (DNase) digestion accompanied with a serious of organic or inorganic pretreatment. This protocol is easy to operate and ensures high efficiency for the decellularization. The TEM showed favorable cytocompatibility for TSCC cells under static or stirred culture conditions, which enables the construction of the TSCC model. A self-made bioreactor was also used for the persistent stirred condition for cell culture. Reconstructed TSCC using TEM showed the characteristics and properties resembling clinical TSCC histopathology, suggesting the potential in TSCC research.

Fabrication of Tongue Extracellular Matrix and Reconstitution of Tongue Squamous Cell Carcinoma In Vitro

A method is shown here for the preparation of the tongue extracellular matrix (TEM) with efficient decellularization. The TEM can be used as functional scaffolds for the reconstruction of a tongue squamous cell carcinoma (TSCC) model under static or stirred culture conditions.

In order to construct an effective and realistic model for tongue squamous cell carcinoma (TSCC) in vitro, the methods were created to produce ...

Extraction of Extracellular Vesicles from Whole Tissue

Circulating and interstitial small membrane-bound extracellular vesicles (EVs) represent promising targets for the development of novel diagnostic or prognostic biomarker assays, and likely serve as important players in the progression of a vast spectrum of diseases. Current research is focused on the characterization of vesicles secreted from multiple cell and tissue types in order to better understand the role of EVs in the pathogenesis of conditions including neurodegeneration, inflammation, and cancer. However, globally consistent and reproducible techniques to isolate and purify vesicles remain in progress. Moreover, methods for extraction of EVs from solid tissue ex vivo are scarcely described. Here, we provide a detailed protocol for extracting small EVs of interest from whole fresh or frozen tissues, including brain and tumor specimens, for further characterization. We demonstrate the adaptability of this method for multiple downstream analyses, including electron microscopy and immunophenotypic characterization of vesicles, as well as quantitative mass spectrometry of EV proteins.

Here, we provide a detailed protocol to isolate small extracellular vesicles (EVs) from whole tissues, including brain and tumor specimens. This method offers a reproducible technique to extract EVs from solid tissue for further downstream analyses.

Circulating and interstitial small membrane-bound extracellular vesicles (EVs) represent promising targets for the development of novel diagnostic or ...

Three-Dimensional Bone Extracellular Matrix Model for Osteosarcoma

Osteosarcoma (OS) is the most common and a highly aggressive primary bone tumor. It is characterized with anatomic and histologic variations along with diagnostic or prognostic difficulties. OS comprises genotypically and phenotypically heterogeneous cancer cells. Bone microenvironment elements are proved to account for tumor heterogeneity and disease progression. Bone extracellular matrix (BEM) retains the microstructural matrices and biochemical components of native extracellular matrix. This tissue-specific niche provides a favorable and long-term scaffold for OS cell seeding and proliferation. This article provides a protocol for the preparation of BEM model and its further experimental application. OS cells can grow and differentiate into multiple phenotypes consistent with the histopathological complexity of OS clinical specimens. The model also allows visualization of diverse morphologies and their association with genetic alterations and underlying regulatory mechanisms. As homologous to human OS, this BEM-OS model can be developed and applied to the pathology and clinical research of OS.

The bone extracellular matrix (BEM) model for osteosarcoma (OS) is well established and shown here. It can be used as a suitable scaffold for mimicking primary tumor growth in vitro and providing an ideal model for studying the histologic and cytogenic heterogeneity of OS.

Osteosarcoma (OS) is the most common and a highly aggressive primary bone tumor. It is characterized with anatomic and histologic variations along with ...

Cell Cycle-specific Measurement of &#947;H2AX and Apoptosis After Genotoxic Stress by Flow Cytometry

The presented method or slightly modified versions have been devised to study specific treatment responses and side effects of various anti-cancer treatments as used in clinical oncology. It enables a quantitative and longitudinal analysis of the DNA damage response after genotoxic stress, as induced by radiotherapy and a multitude of anti-cancer drugs. The method covers all stages of the DNA damage response, providing endpoints for induction and repair of DNA double-strand breaks (DSBs), cell cycle arrest and cell death by apoptosis in case of repair failure. Combining these measurements provides information about cell cycle-dependent treatment effects and thus allows an in-depth study of the interplay between cellular proliferation and coping mechanisms against DNA damage. As the effect of many cancer therapeutics including chemotherapeutic agents and ionizing radiation is limited to or strongly varies according to specific cell cycle phases, correlative analyses rely on a robust and feasible method to assess the treatment effects on the DNA in a cell cycle-specific manner. This is not possible with single-endpoint assays and an important advantage of the presented method. The method is not restricted to any particular cell line and has been thoroughly tested in a multitude of tumor and normal tissue cell lines. It can be widely applied as a comprehensive genotoxicity assay in many fields of oncology besides radio-oncology, including environmental risk factor assessment, drug screening and evaluation of genetic instability in tumor cells.

The presented method combines the quantitative analysis of DNA double-strand breaks (DSBs), cell cycle distribution and apoptosis to enable cell cycle-specific evaluation of DSB induction and repair as well as the consequences of repair failure.

The presented method or slightly modified versions have been devised to study specific treatment responses and side effects of various anti-cancer ...

Characterization of the Effects of Migrastatic Inhibitors on 3D Tumor Spheroid Invasion by High-resolution Confocal Microscopy

Drug discovery and development in cancer research is increasingly being based on drug screens in a 3D format. Novel inhibitors targeting the migratory and invasive potential of cancer cells, and consequently the metastatic spread of disease, are being discovered and considered as complementary treatments in highly invasive cancers such as gliomas. Thus, generating data enabling the detailed analyses of cells in a 3D environment following the addition of a drug is required. The methodology described here, combining spheroid invasion assays with high-resolution image capture and data analysis by confocal laser scanning microscopy (CLSM), enabled detailed characterization of the effects of the potential anti-migratory inhibitor MI-192 on glioma cells. Spheroids were generated from cell lines for invasion assays in low adherent 96-well plates and then prepared for CLSM analysis. The described workflow was preferred over other commonly used spheroid-generating techniques due to both ease and reproducibility. This, combined with the enhanced image resolution attained by confocal microscopy compared to conventional wide-field approaches, allowed the identification and analysis of distinct morphological changes in migratory cells in a 3D environment following treatment with the migrastatic drug MI-192.

The effects of migrastatic inhibitors on glioma cancer cell migration in three-dimensional (3D) invasion assays using a histone deacetylase (HDAC) inhibitor are characterized by high-resolution confocal microscopy.

Drug discovery and development in cancer research is increasingly being based on drug screens in a 3D format. Novel inhibitors targeting the migratory and ...

Studying Triple Negative Breast Cancer Using Orthotopic Breast Cancer Model

Triple-negative breast cancer (TNBC) is an aggressive breast cancer subtype with limited therapeutic options. When compared to patients with less aggressive breast tumors, the 5-year survival rate of TNBC patients is 77% due to their characteristic drug-resistant phenotype and metastatic burden. Toward this end, murine models have been established aimed at identifying novel therapeutic strategies limiting TNBC tumor growth and metastatic spread. This work describes a practical guide for the TNBC orthotopic model where MDA-MB-231 breast cancer cells suspended in a basement membrane matrix are implanted in the fourth mammary fat pad, which closely mimics the cancer cell behavior in humans. Measurement of tumors by caliper, lung metastasis assessment via in vivo and ex vivo imaging, and molecular detection are discussed. This model provides an excellent platform to study therapeutic efficacy and is especially suitable for the study of the interaction between the primary tumor and distal metastatic sites.

This work presets an advanced protocol to accurately assess tumor loading by detection of green fluorescent protein and bioluminescence signals as well as the integration of quantitative molecular detection technique.

Triple-negative breast cancer (TNBC) is an aggressive breast cancer subtype with limited therapeutic options. When compared to patients with less ...

Studying the Effects of Tumor-Secreted Paracrine Ligands on Macrophage Activation using Co-Culture with Permeable Membrane Supports

Tumor-derived paracrine signaling is an overlooked component of local immunosuppression and can lead to a permissive environment for continued cancer growth and metastasis. Paracrine signals can involve cell-cell contact between different cell types, such as PD-L1 expressed on the surface of tumors interacting directly with PD-1 on the surface of T cells, or the secretion of ligands by a tumor cell to affect an immune cell. Here we describe a co-culture method to interrogate the effects of tumor-secreted ligands on immune cell (macrophage) activation. This straightforward procedure utilizes commercially available 0.4 &#181;m polycarbonate membrane permeable supports and standard tissue culture plates. In the process described, macrophages are cultured in the upper chamber and tumor cells in the lower chamber. The presence of the 0.4 &#181;m barrier allows for the study of intercellular signaling without the confounding variable of physical contact, because the two cell types share the same medium and exposure to paracrine ligands. This approach can be combined with others, such as genetic alterations of the macrophage (e.g., isolation from genetic knock-out mice) or tumor (e.g., CRISPR-mediated alterations) to study the role of specific secreted factors and receptors. The approach also lends itself to standard molecular biological analyses such as quantitative reverse transcription polymerase chain reaction (qRT-PCR) or Western blot analysis, without the need for flow sorting to separate the two cell populations. Enzyme-linked immunosorbent assays (ELISAs) can similarly be utilized to measure secreted ligands to better understand the dynamic interaction of cell signaling in the multiple cell type context. Duration of co-culture can also be varied for the study of temporally regulated events. This co-culture method is a robust tool that facilitates the study of tumor-secreted signals in the immune context.

Here, we present a method using permeable membrane supports to facilitate the study of non-contact paracrine signaling used by tumor cells to suppress the immune response. The system is amenable to studying the role of tumor-secreted factors in dampening macrophage activation.

Tumor-derived paracrine signaling is an overlooked component of local immunosuppression and can lead to a permissive environment for continued cancer ...

Studying Normal Tissue Radiation Effects using Extracellular Matrix Hydrogels

Summary

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Chapters in this video

Analyzing the Communication Between Monocytes and Primary Breast Cancer Cells in an Extracellular Matrix Extract (ECME)-based Three-dimensional System

PET and MRI Guided Irradiation of a Glioblastoma Rat Model Using a Micro-irradiator

Imaging Approaches to Assessments of Toxicological Oxidative Stress Using Genetically-encoded Fluorogenic Sensors

Fabrication of Tongue Extracellular Matrix and Reconstitution of Tongue Squamous Cell Carcinoma In Vitro

Extraction of Extracellular Vesicles from Whole Tissue

Three-Dimensional Bone Extracellular Matrix Model for Osteosarcoma

Cell Cycle-specific Measurement of γH2AX and Apoptosis After Genotoxic Stress by Flow Cytometry

Characterization of the Effects of Migrastatic Inhibitors on 3D Tumor Spheroid Invasion by High-resolution Confocal Microscopy

Studying Triple Negative Breast Cancer Using Orthotopic Breast Cancer Model

Studying the Effects of Tumor-Secreted Paracrine Ligands on Macrophage Activation using Co-Culture with Permeable Membrane Supports