Ms. Audrey Thomas-Gahring is acknowledged for her GC/MS work. We would also like to thank Massimo Marzorati for editing the manuscript.

1. Materials and Preparations
NOTE: The defined medium is purchased as a powder (see Table of Materials). The composition of the defined medium in g/L is the following: Arabinogalactan (1.2), Pectin (2.0), Xylan (0.5), Glucose (0.4), Yeast extract (3.0), Special peptone (1.0), Mucin (2.0), L-cysteine-HCl (0.2).
<ol>
	<li>Prepare the defined medium
	<ol>
		<li>Fill a 4 L flask with 2 L of double distilled, deionized water.</li>
		<li>Add 29.2 g of defined medium powder to the water and mix unit complete homogenized.</li>
		<li>Add a magnetic stirrer and loosely screw on the lid.</li>
		<li>Autoclave at 121 &#176;C for 30 min.</li>
		<li>After cooling, store the prepared defined medium in a refrigerator at 4 &#176;C with constant agitation (magnetic stirrer).</li>
	</ol>
	</li>
	<li>Prepare pancreatic juice
	<ol>
		<li>Autoclave 2L of distilled, deionized water in a 5L vessel with a stirrer bar added, at 121 &#176;C for 30 min.</li>
		<li>After autoclaving, allow the water to cool to room temperature.</li>
		<li>Add 12.5 g NaHCO3, 6 g bile, and 0.9 g pancreatin. Place on magnetic stirrer to mix. 
		NOTE: The pancreatic juice should be prepared fresh every 2-3 days and refrigerated with agitation after it made.</li>
	</ol>
	</li>
	<li>Phosphate buffer
	<ol>
		<li>Place a 1 L flask containing a magnetic stirrer bar on a magnetic stirrer. Add 0.5 L of distilled, deionized water and turn on agitation.</li>
		<li>Next, add 6.8 g KH2PO4, 8.8 g K2HPO4, and 0.1 g sodium thioglycolate. Top off with water to a final volume of 1 L.</li>
		<li>After the components have completely dissolved, adjust the pH to 7.0 using NaOH.</li>
		<li>Pour the buffer into a 2 L vessel with a screw cap lid and autoclave at 121 &#176;C for 30 min. Ensure that the lid is screwed on loosely and not closed tightly.</li>
		<li>After removing from the autoclave, allow the solution to cool to the room temperature. Store the buffer at room temperature for up to 2 weeks.</li>
	</ol>
	</li>
	<li>Prepare Mucin agar carriers
	<ol>
		<li>Autoclave plastic mucin carriers (see Table of Materials), forceps, plastic mesh (tubular shape), and zip ties at 131 &#176;C for 30 min.</li>
		<li>Autoclave 300 mL of double distilled, de-ionized water at 131 &#176;C for 30 min. Store at room temperature until use.</li>
		<li>In a biosafety cabinet with the laminar flow, fill sterile Petri dishes with the plastic mucin carriers using sterile forceps. 
		NOTE: The mucin carriers are hollow plastic circles that are filled with mucin agar. Once filled, the lid can be placed on top and these can be set aside.</li>
		<li>To prepare mucin agar, add 3 g of bacterial agar and 15 g of porcine mucin to the 300 mL of autoclaved water.</li>
		<li>In a fume hood, boil this solution, then remove from the heat source and allow to cool for 1 min. Repeat the boiling and cooling for a total of 3 times.</li>
		<li>Once the glass vessel containing the mucin agar solution is cool enough to touch, move it into the biosafety cabinet with laminar flow.</li>
		<li>In the biosafety cabinet with the laminar flow, pour the liquid mucin agar over the plastic mucin carriers that were placed in the Petri dish. Ensure that all the carriers are filled completely with mucin agar.</li>
		<li>Place the lid back on the Petri dish and allow it to cool under laminar flow.</li>
		<li>To assemble the mucin carriers, take the autoclaved plastic netting and insert mucin carriers containing the solidified mucin agar from the Petri dish using sterile forceps.</li>
		<li>Use zip ties to close the ends of the mesh. In this way the net functions to contain the mucin carriers filled with mucin agar. Store at 4 &#176;C until use.</li>
	</ol>
	</li>
</ol>
2. Set up, Inoculation, and Running of the System
<ol>
	<li>Set up of the culturing system 
	NOTE: The Twin Simulator of the Human Intestinal Microbial Ecosystem was used for this study.
	<ol>
		<li>Place 10 bioreactors containing stirrer bars on magnetic stirrers and turn on agitation. 
		NOTE: Each Unit will consist of 5 bioreactors, so 2 units will require 10 bioreactors.</li>
		<li>Connect 5 bioreactors to each other using silicon tubing in sequence by way of the peristaltic pump (Figure 1). Use the pumps to transverse the fluid contents from one bioreactor to the next.</li>
		<li>Label the bioreactors in the following sequence: Stomach, small intestine, ascending colon, transverse colon, descending colon to represent the order of the gastrointestinal tract. 
		NOTE: The bioreactors can all be identical, or the stomach and small intestine bioreactors can be smaller compared to the colon bioreactors since they will hold less volume. Setting up the bioreactors in this way means that the ascending colon region receives nutrients from the small intestine, the transverse colon region receives nutrients from the ascending colon region, and the descending colon region receives nutrients from the transverse region. In this way, the system is mimicking the sequential movement of nutrients through the gastrointestinal tract.</li>
		<li>Place the defined medium and pancreatic juice in a refrigerator at 4 &#176;C with constant agitation using magnetic stirrers. Using silicon tubing, connect the defined medium to the stomach bioreactor via a peristaltic pump and connect the pancreatic juice to the small intestine via a peristaltic pump.</li>
		<li>Add solidifying agent to a urine drainage bag. Connect the descending colon to the urine drainage bag by way of the peristaltic pump. Dispose of the solidified as biological waste during the experiment.</li>
		<li>Using silicone tubing, connect the water jacket of each vessel in order, and connect each end to the circulating water bath. Fill the circulating water bath with distilled water (this amount will vary depending on the type of circulating water bath used) and set to 37 &#176;C.</li>
		<li>Using silicon tubing, connect the acid and the base to the lid of each vessel by way of peristaltic pumps. The recommended concentrations are 0.5M HCl and 0.5M NaOH.</li>
		<li>Insert the pH probes to each vessel through the designated port in the lid. This port is designed to hold the pH probe and screw into the lid (Figure 1). Set the pH for each region as follows: Stomach, pH = 2; Small intestine, pH = 6.7-6.9; Ascending colon, pH = 5.6-5.9; Transverse colon, pH = 6.15-6.4; Descending colon, pH = 6.6-6.9. 
		NOTE: During the initial phase, the community will differentiate based on the difference in region pH values. However, once the feeding cycles begin, the communities will further mature based on the differences in nutrient input.</li>
		<li>Connect a nitrogen line using silicon tubing to the lid of each vessel using the designated port. The nitrogen line should flow in the following sequence: stomach, small intestine, ascending colon, transverse colon, descending colon. Ensure each unit has its own flush to avoid cross contamination. 
		NOTE: Nitrogen is used to removed oxygen from the system and maintains anaerobiosis during the experiment.</li>
		<li>Insert a metal sample tube through the lid of each bioreactor using the designated port. The metal tube extends down into the bioreactor and is used to collect fluid samples.</li>
		<li>Add a small piece of silicon tubing to the top end of the sample tube and connect a Luer Lock to allow for the use of a syringe during sampling.</li>
	</ol>
	</li>
	<li>Inoculation of the system
	<ol>
		<li>Fill the colon regions with the defined medium using the following volumes: 500 mL in the ascending colon, 800 mL in the transverse colon, and 600 mL in the descending colon.</li>
		<li>Add mucin agar carriers to the colon regions, in an amount proportional to the volume of the reactor. For this experiment, 60 carriers per reactor were used.</li>
		<li>Remove any excess defined medium using the sample tube and a syringe since adding the carriers raises the level of fluid in the bioreactor.</li>
		<li>Turn on the pH probes to adjust the pH in each reactor using the computer.</li>
		<li>Turn on the nitrogen flush for 20 min. 
		NOTE: The fecal homogenate used for inoculation is purchased as 10% feces homogenized in 10% glycerol solution (see Table of Materials).The fecal sample is randomly selected from a pool of donors with the following criteria: An American consuming a typical western diet (not vegan or vegetarian), between 21-45 years of age, antibiotic free for at least 1 year, with an average Body Mass Index (18.5-24.9). For this protocol, only a single donor is used. However, this can be altered due to experimental design.</li>
		<li>Prior to inoculation, thaw the fecal homogenate following the manufacturer&#8217;s guidelines.</li>
		<li>Inoculate each colon reactor by adding the fecal homogenate in an amount equal to 5% of the reactor volume through the sample port, using a 60 mL syringe (25 mL for the ascending colon, 40 mL for the transverse colon, 30 mL for the descending colon).</li>
		<li>Grow the cultures in the bioreactors overnight with pH control.</li>
		<li>After the overnight growth, harvest samples of the luminal fluid from each colon region as detailed below in section 3. After sampling is completed, turn on the computer program to begin feeding.</li>
	</ol>
	</li>
	<li>Daily feeding cycles 
	NOTE: The daily feeding cycles are computer automated.
	<ol>
		<li>Three times a day, pump 140 mL of the defined medium into the stomach bioreactor and allowed to incubate for 1 h.</li>
		<li>After 1 h incubation, pump the contents of the stomach into the small intestine. At the same time, pump 60 mL of pancreatic juice into the small intestine.</li>
		<li>In the small intestine, turn on pH adjustment to pH 6.7-6.9, and allow mixture to incubate for 90 mins. During incubation, turn on the nitrogen flush for each system for a total of 10 min each.</li>
		<li>After the 90 min incubation, turn on the pumps from the small intestine to the ascending colon, the ascending colon to the transverse colon, the transverse colon to the descending colon, and the descending colon to the waste simultaneously.</li>
	</ol>
	</li>
	<li>Experimental timeline 
	NOTE: The in vitro system used here is operated in a continuous fashion for approximately 8 weeks in length. Below is a recommended experimental timeline, however, this can be modified depending on the requirements and objective of the experiment.
	<ol>
		<li>Set up system following steps listed in step 2.1.</li>
		<li>Harvest samples from overnight growth following the protocol below. Begin the daily cycles, feeding three times a day following the protocol above.</li>
		<li>Run the experiment for a total of 2 weeks to allow the bacteria to stabilize. 
		NOTE: Running the system means that the pH is maintained, the daily feeding cycles are followed three times a day, and the mucosal carriers are changed 2-3 times each week.</li>
		<li>After stabilization, run the experiment for 2 weeks as the control period.</li>
		<li>After the control period, use the system to test different components or variables, known as the treatment period.</li>
		<li>After the treatment period, stop adding the components or variables and return the conditions to normal. This is considered the wash-out period and is used to determine whether or not changes to the community are permanent.</li>
	</ol>
	</li>
</ol>
3. Harvesting Samples from the System
NOTE: During the experiment, samples can be harvested from the luminal or mucosal phase of any region at any time, following the below guidelines.
<ol>
	<li>Harvesting luminal samples
	<ol>
		<li>Harvest luminal fluid samples through the sample port that extends into the middle of the culture liquid. Begin by cleaning the Luer Lock on the sample port with either 70% ethanol or the small alcohol pad and allow to dry.</li>
		<li>Ensure that all air is removed from a 30 mL syringe and connect it to the sample port. Draw up and down 3-5 times to ensure proper mixing of the vessel components. After, draw a total of 30 mL of culture.</li>
		<li>Aliquot the luminal samples as needed into sterile falcon tubes and store on ice. After sampling is completed transfer these to a -80 &#176;C freezer.</li>
		<li>For SCFA analysis, 15 mL of the luminal sample is spun at 4 &#176;C, 5,000 x g for 10 min. The supernatant is syringe filtered using a PES 0.2 &#956;M filter. Store the filtered supernatant at -80 &#176;C.
		<ol>
			<li>For DNA analysis, spin 1 mL of luminal fluid at 4 &#176;C, 5,000 x g for 10 min. Discard the supernatant and store the pellet at -80 &#176;C.</li>
			<li>As a backup, store 10 mL of luminal fluid at -80 &#176;C for most experiments.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Harvesting mucosal samples 
	NOTE: Change mucosal carriers every 2-3 days.
	<ol>
		<li>Remove and bring out the prepared mucin carriers from the refrigerator.</li>
		<li>Turn on the nitrogen flush and open the reactor lid quickly. Remove 50% of mucin carriers and add new ones.</li>
		<li>Seal the lid quickly and keep the nitrogen flush on for another 20 min.</li>
		<li>For DNA extraction, aliquot 0.25-0.5 g of agar into a 2 mL tube and stored at -80 &#176;C until needed.</li>
	</ol>
	</li>
</ol>
4. DNA Extraction, Sequencing, and Analysis
NOTE: DNA is extracted using the CTAB DNA extraction method with physical homogenization in a fume hood29. Following extraction, a spectrophotometer is used to quantify the amount of DNA in each sample.
<ol>
	<li>Prepare CTAB buffer
	<ol>
		<li>Dissolve 4.2 g K2HPO4 and 4.09 g NaCl in 200 mL of deionized, distilled water, then autoclave at 121 &#176;C for 30 min.</li>
		<li>Allow the solution to cool to room temperature.</li>
		<li>Add 10 g of Hexadecyltrimethylammonium bromide (CTAB) and heat to 60 &#176;C with agitation.</li>
		<li>After all the CTAB is dissolved, remove the solution from heat and cool to room temperature.</li>
	</ol>
	</li>
	<li>Prepare PEG-6000 solution
	<ol>
		<li>Dissolve 300 g of polyethylene glycol 6,000 and 93.5 g NaCl in 1 L of deionized, distilled water.</li>
		<li>After all particles are dissolved, autoclave the solution at 121 &#176;C for 30 min.</li>
		<li>After autoclaving, cool the solution to room temperature before use.</li>
	</ol>
	</li>
	<li>Perform DNA extraction
	<ol>
		<li>Add 500 &#956;L of CTAB buffer and 500 &#956;L of Phenol-Chloroform-Isoamyl alcohol to the 2 mL sample tube and vortex to homogenize.</li>
		<li>Transfer the entire contents of the sample tube to a 0.1 mm physical homogenizing tube (see Table of Materials).</li>
		<li>Homogenize the tubes for 20 s at the highest setting two times, with a 20 s pause in between, using a physical homogenizer (see Table of Materials).</li>
		<li>Centrifuge the homogenized tubes at 3,000 x g for 5 mins.</li>
		<li>Transfer 300 &#956;L of the supernatant to a clean, 1.7 mL tube and add 500 &#956;L of CTAB buffer to the original tube.</li>
		<li>Homogenize this tube again using the methods in steps 4.3.2-4.3.3 and centrifuge at 3,000 x g for 5 min.</li>
		<li>After centrifugation, transfer another 300 &#956;L of supernatant to the new tube which now contains 600 &#956;L.</li>
		<li>Add a total of 600 &#956;L Chloroform-Isoamyl alcohol to the tube containing 600 &#956;L of the supernatant.</li>
		<li>Invert the tubes to mix, and then centrifuge briefly. Transfer 500 &#956;L of the upper phase to a clean 1.7 mL tube. Ensure only to take the top phase.</li>
		<li>Next, add 1000 &#956;L of Peg-6000 solution, invert the tubes to mix, and then incubate at room temperature for 2 h.</li>
		<li>After incubation, centrifuge the tubes at 18,200 x g, 4 &#176;C, for 10 min.</li>
		<li>Discard the supernatant and clean the pellet with ice cold 70% ethanol.</li>
		<li>Centrifuge the tubes again at 18,200 x g, 4 &#176;C, for 10 min. Discard the supernatant and allow the pellet to dry in the biosafety cabinet under laminar flow.</li>
		<li>After the pellet is dry, add 75 &#956;L of RNAse free, DNAse free, sterile water to each tube.</li>
		<li>Quantify the resuspended DNA by a spectrophotometer and then send it out for 16S rRNA gene sequencing.</li>
	</ol>
	</li>
</ol>
5. Short Chain Fatty Acid (SCFA) Detection and Analysis
<ol>
	<li>Thaw the frozen samples at 40&#176;C for 30 min and extract short chain fatty acids using diethyl ether in a 2:1 (v:v) ratio.</li>
	<li>Transfer the organic phase to a GC/MS (see Table of Materials) equipped with a column, 30 m, 0.25 mm ID, 0.25 &#181;m, (see Table of Materials) where 10 &#181;L is injected.</li>
	<li>Set the injection port temperature and initial column temperature to 260 &#176;C and 125 &#176;C respectively. Hold the initial temperature for 1 min, and then increase this to 250 &#176;C, over 11.5 min with a column flow rate of 1 mL/min. 
	NOTE: The MS temperature for the ion source is 220&#176;C and 250&#176;C for the interface.</li>
	<li>Quantify the amount of each SCFA using standard curves and 2-methylhexanoic acid as the internal standard.</li>
</ol>

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href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemostat+culture+systems+support+diverse+bacteriophage+communities+from+human+feces.">Chemostat culture systems support diverse bacteriophage communities from human feces.</a> Microbiome. 9 (3), 58 (2015).</li><li>Sousa, T., Paterson, R., Moore, V., Carlsson, A., Abrahamsson, B., Basit, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+gastrointestinal+microbiota+as+a+site+for+the+biotransformation+of+drugs.">The gastrointestinal microbiota as a site for the biotransformation of drugs.</a> International Journal of Pharmaceutics. 363 (1-2), 1-25 (2008).</li><li>Pferschy-Wenzig, E., Koskinen, K., Moissl-Eichinger, C., Bauer, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Combined+LC-MS+Metabolomics-+and+16S+rRNA+Sequencing+Platform+to+Assess+Interactions+between+Herbal+Medicinal+Products+and+Human+Gut+Bacteria+in+Vitro:+a+Pilot+Study+on+Willow+Bark+Extract.">A Combined LC-MS Metabolomics- and 16S rRNA Sequencing Platform to Assess Interactions between Herbal Medicinal Products and Human Gut Bacteria in Vitro: a Pilot Study on Willow Bark Extract.</a> Frontiers in Pharmacology. 8 (893), (2017).</li><li>Cueva, C., 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=In+vitro+fermentation+of+grape+seed+flavan-3-ol+fractions+by+human+faecal+microbiota:+changes+in+microbial+groups+and+phenolic+metabolites.">In vitro fermentation of grape seed flavan-3-ol fractions by human faecal microbiota: changes in microbial groups and phenolic metabolites.</a> FEMS Microbiology Ecology. 83 (3), 792-805 (2013).</li><li>Molly, K., Van de Woestyne, M., Verstraete, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+a+5-step+multi-chamber+reactor+as+a+simulation+of+the+human+intestinal+microbial+ecosystem.">Development of a 5-step multi-chamber reactor as a simulation of the human intestinal microbial ecosystem.</a> Applied Microbiology and Biotechnology. 39 (2), 254-258 (1993).</li><li>Wang, M., 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=Apigenin+Impacts+the+Growth+of+the+Gut+Microbiota+and+Alters+the+Gene+Expression+of+Enterococcus.">Apigenin Impacts the Growth of the Gut Microbiota and Alters the Gene Expression of Enterococcus.</a> Molecules. 22 (8), (2017).</li><li>Caporaso, J., 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=QIIME+allows+analysis+of+high-throughput+community+sequencing+data.">QIIME allows analysis of high-throughput community sequencing data.</a> Nature Methods. 7 (5), 335-336 (2010).</li><li>Edgar, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Search+and+clustering+orders+of+magnitude+faster+than+BLAST.">Search and clustering orders of magnitude faster than BLAST.</a> Bioinformatics. 26 (19), 2460-2461 (2010).</li><li>Cole, J., 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=Ribosomal+Database+Project:+data+and+tools+for+high+throughput+rRNA+analysis.">Ribosomal Database Project: data and tools for high throughput rRNA analysis.</a> Nucleic Acids Research. 42 (database issue), 633-642 (2014).</li><li>McDonald, D., 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=An+improved+Greengenes+taxonomy+with+explicit+ranks+for+ecological+and+evolutionary+analyses+of+bacteria+and+archaea.">An improved Greengenes taxonomy with explicit ranks for ecological and evolutionary analyses of bacteria and archaea.</a> The ISME Journal. 6 (3), 610-618 (2012).</li><li>Liu, L. S., 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=Establishing+a+mucosal+gut+microbial+community+in+vitro+using+an+artificial+simulator.">Establishing a mucosal gut microbial community in vitro using an artificial simulator.</a> PLoS ONE. 13 (7), e0197692 (2018).</li><li>Van de Wiele, T., Van den Abbeele, P., Ossieur, W., Possemiers, S., Marzorati, M., Verhoeckx, K., 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=Chapter+27:+The+Simulator+of+the+Human+Intestinal+Microbial+Ecosystem+(SHIME+&#174;).">Chapter 27: The Simulator of the Human Intestinal Microbial Ecosystem (SHIME &#174;).</a> The Impact of Food Bio-Actives on Gut Health. , (2015).</li><li>Van den Abbeele, P., 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=Butyrate-producing+Clostridium+cluster+XIVa+species+specifically+colonize+mucins+in+an+in+vitro+gut+model.">Butyrate-producing Clostridium cluster XIVa species specifically colonize mucins in an in vitro gut model.</a> The ISME Journal. 7 (5), 949-961 (2013).</li><li>Possemiers, S., Verth&#233;, K., Uyttendaele, S., Verstraete, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PCR-DGGE-based+quantification+of+stability+of+the+microbial+community+in+a+simulator+of+the+human+intestinal+microbial+ecosystem.">PCR-DGGE-based quantification of stability of the microbial community in a simulator of the human intestinal microbial ecosystem.</a> FEMS Microbiology Ecology. 49 (3), 495-507 (2004).</li><li>Tan, J., McKenzie, C., Potamitis, M., Thorburn, A., Mackay, C., Macia, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+short-chain+fatty+acids+in+health+and+disease.">The role of short-chain fatty acids in health and disease.</a> Advances in Immunology. 121, 91-119 (2014).</li><li>Yang, J., Kweon, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+gut+microbiota:+a+key+regulator+of+metabolic+diseases.">The gut microbiota: a key regulator of metabolic diseases.</a> BMB Reports. 49 (10), 536-541 (2016).</li><li>Sonnenburg, J., B&#228;ckhed, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diet-microbiota+interactions+as+moderators+of+human+metabolism.">Diet-microbiota interactions as moderators of human metabolism.</a> Nature. 535 (7610), 56-64 (2016).</li></ol>

The authors have no competing financial interests. Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U.S. Department of Agriculture. USDA is an equal opportunity provider and employer.

In vitro culturing systems have been developed to study the gut microbiota of the large intestine. They use apparatuses designed to simulate the physiological conditions of the gastro intestinal tract, promoting the growth of a mature gut microbial community for each region of the colon33. While the concept is logical and comprehensible, the actual running of in vitro culturing systems to study the gut microbiota requires precision and an understanding of what is required and expected to produce r...

The gut microbiota is a community of micro-organisms that reside in the human gastrointestinal tract (GIT).&#160; This community reaches maximum concentration in the colon, which is estimated to hold 1013-1014 bacteria, from 500-1,000 species, that live in symbiosis with the colon milieu1,2. The composition and functionality of the gut microbiota change spatially along the GIT, forming region specific communities, with the most diversity found distally2,3,4,5. For each anatomical region, separate microbial communities reside in the lumen and on the mucosal lining6. The lumen community has more direct access to nutrients as substrates move through the luminal compartment7. Despite this, some bacteria reside preferentially in the mucus layer, utilizing mucin produced by the colon cells as an energy source1,5,8. The difference in microenvironments between the luminal and mucosal phases results in divergence and the development of phase specific communities. Together, these communities provide metabolic functions, such as nutrient metabolism and the production of vitamins, and immunological functions, such as preventing the colonization of human pathogens1,3,9. The gut microbiota also works functionally in conjugation with the human colon cells3.
As an important part of the human GIT, it is not surprising that the gut microbiota is known to contribute to both host health and disease status3,9,10,11,12. A shift in the gut microbial population has been associated with multiple human diseases, including GIT disorders like intestinal bowel disease (IBD) and intestinal bowel syndrome (IBS), but also other diseases, such as obesity, circulatory disease, and autism3,9,10,11,12.&#160; Metabolites produced from the gut microbiota have a global effect, reaching locations far from the gut12,13. For example, the gut-brain axis is associated with mental disorders like anxiety and depression14. Therefore, studying the gut microbiota is important to multiple fields of research, and is applicable to many diseases, even those not often associated with the GIT.
While it is widely acknowledged that studying the gut microbiota is important, it is a complicated endeavor. Multiple animal models are available, from small animals like zebrafish, rats, and mice, to larger ones like monkeys and pigs15-19. However, the application of these animals in terms of the human gut microbiota is not straightforward, since these animals have a unique bacterial community that has evolved based on environment and diet, and they are anatomically distinct from humans20,21. The use of human subjects removes the question of relevance yet introduces another set of challenges. Human studies are expensive, time consuming, and are ethically constrained11. Moreover, confounding factors influence the gut microbiota in human studies, including age or developmental stage, environment, diet, medication, and genetic factors2,4,22.&#160; There are also restrictions on what can be tested in humans, and which type of samples can be harvested at what times4.&#160;
One critical disadvantage of using an in vivo system to study the gut microbiota is the presence of mammalian components. The gut microbiota and human cells interact with one another, and in an in vivo setting, it is impossible to distinguish the two. The metabolites produced by the gut microbiota are taken in by the colon cells, so measurements cannot be calculated with precision. Therefore, any mechanistic study must be limited to end-point measurements11. Another major disadvantage for in vivo studies is the inability to harvest samples from the different regions of the GIT longitudinally23. This does not allow for the assessment of changes that may occur in the microenvironments of the colon over time12.&#160; Many in vivo studies, including human studies, rely on analysis of fecal samples to detect changes to the gut microbiota12. While this is informative, it does not provide data on the gut microbiota across the GIT and does not differentiate between the luminal and mucosal communities5,6,7,8.
For the gut microbiota, the application of an in vitro method is required to study the dynamics of the bacterial community, without interference from the mammalian components. Using an in vitro method allows for the tight control of environmental conditions10, testing of multiple parameters simultaneously, and the ability to sample longitudinally, and in large volumes11. Since an in vitro method utilizes a mechanical device and not a host, no considerations are needed for age, environment, diet, or genetic background. These systems can be used to test either the entire gut microbiota community, only selected organisms, or even single strains. Importantly, in vitro results are reproducible, yet retain a level of diversity comparable to in vivo studies11,22.
Depending on the hypothesis in question and the desired results, in vitro studies can be performed in numerous ways.&#160; They can utilize single-vessel systems and simple methods, such as incubating samples with fecal homogenate24 or performing single batch cultures over the course of 24-48 h25. They can also be accomplished using single-vessel systems and more complex methods, such as using a chemostat system to produce a stable gut microbial community11. However, the use of a single reactor can over-simplify the microbiota12 since it only represents one section of the colon, even though the colon is composed of the ascending, transverse, and descending regions.
In order to study the gut microbiota community that develops in the different regions of the colon (the ascending, transverse, and descending regions), a complex, multi-stage system can be employed. In these systems, multiple vessels are set up to mimic the different regions of the colon, so the gut microbiota of the ascending, transverse, and descending regions are cultivated independently. These vessels are connected, using pumps to move substrates in sequence, from the ascending to the transverse to the descending colon regions, mimicking the flow of nutrients through the GIT.
The objective of this study was to demonstrate how a 5-stage in vitro culture system (see Table of Materials) can be used to cultivate the gut microbiota community, and to demonstrate community dynamics in terms of stability and composition. In this system, one vessel represents the stomach and one represent the small intestine. The colon is divided into three regions (ascending, transverse, descending), with one vessel representing each region26. In this experimental setup, two complete systems were run in parallel, with Unit 1 containing mucin carriers to represents the mucosal surface and Unit 2 containing no mucin carriers. The communities that developed in the luminal and mucosal phases of each region were compared to each other, and to the fecal inoculum over time using 16S rRNA gene sequencing and SCFA analysis. The results presented demonstrate the type of community, both in terms of composition and functionality, which can be produced from this type of in vitro system.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>TWINSHIME</td>
			<td>Prodigest</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>defined medium (Adult M-SHIME growth medium with starch)</td>
			<td>Prodigest</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Masterflex tubing</td>
			<td>cole Parmer</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Urine Drainage bag</td>
			<td>Bard</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Labsorb</td>
			<td>Sigma-Aldrich</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Fecal sample</td>
			<td>Openbiome</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringes</td>
			<td>Becton Dickson</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Defined medium</td>
			<td>Prodigest</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Oxgall Bile</td>
			<td>Becton Dickson</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Pancreatin</td>
			<td>Sigma-Aldrich</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass ware</td>
			<td>Ace Glass</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Porcine mucin</td>
			<td>Sigma-Aldrich</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Bacteriological agar</td>
			<td>Sigma-Aldrich</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterilization pouches</td>
			<td>VWR</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>BeadBug</td>
			<td>Benchmark Scientific</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Triple-Pure High Impact Zirconium 0.1mm Bead beater tube</td>
			<td>Benchmark Scientific</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>RNAse free, DNAse free, sterile water</td>
			<td>Roche</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Shimadzu QP2010 Ultra GC/MS</td>
			<td>Shimadzu</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Stabilwax-DA column, 30m, 0.25mm ID, 0.25&#181;m</td>
			<td>Restek</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>plastic mucin carriers</td>
			<td>Prodigest</td>
			<td>NA</td>
			<td></td>
		</tr>
	</tbody>
</table>

applying advanced in vitro culturing technology to study the human gut microbiota

The human gut microbiota plays a vital role in both human health and disease. Studying the gut microbiota using an in vivo model, is difficult due to its complex nature, and its diverse association with mammalian components. The goal of this protocol is to culture the gut microbiota in vitro, which allows for the study of the gut microbiota dynamics, without having to consider the contribution of the mammalian milieu. Using in vitro culturing technology, the physiological conditions of the gastro intestinal tract are simulated, including parameters such as pH, temperature, anaerobiosis, and transit time. The intestinal surface of the colon is simulated by adding mucin-coated carriers, creating a mucosal phase, and adding further dimension. The gut microbiota is introduced by inoculating with the human fecal material. Upon inoculation with this complex mixture of bacteria, specific microbes are enriched in the different longitudinal (ascending, transverse and descending colons) and transversal (luminal and mucosal) environments of the in vitro model. It is crucial to allow the system to reach a steady state, in which the community and the metabolites produced remain stable. The experimental results in this manuscript demonstrate how the inoculated gut microbiota community develops into a stable community over time. Once steady state is achieved, the system can be used to analyze bacterial interactions and community functions or to test the effects of any additives on the gut microbiota, such as food, food components, or pharmaceuticals.

The above protocol describes set up, inoculation, and running of a 5-stage in vitro system to study the gut microbiota of the colon. To generate the data presented below, following DNA extraction, 16S rRNA marker gene DNA sequencing of the V1V2 region was performed using the high throughput sequencing (e.g., MiSeq Illumina platform) by the Microbiome Center at the Children&#8217;s Hospital of Philadelphia27. QIIME (Quantitative Insight into Microbial Ecolo...

Watch this Scientific Journal Video about Applying Advanced In Vitro Culturing Technology to Study the Human Gut Microbiota at JoVE.com

Applying Advanced In Vitro Culturing Technology to Study the Human Gut Microbiota

Here, we present a protocol for culturing the gut microbiota of the colon in vitro, using a series of bioreactors that simulate the physiological conditions of the gastro intestinal tract.

The human gut microbiota plays a vital role in both human health and disease. Studying the gut microbiota using an in vivo model, is difficult due to its ...

applying-advanced-vitro-culturing-technology-to-study-human-gut

Research

JoVE Journal

Bioengineering

13.8K Views.  Agricultural Research Service, United States Department of Agriculture. Here, we present a protocol for culturing the gut microbiota of the colon in vitro, using a series of bioreactors that simulate the physiological conditions of the gastro intestinal tract.

Video: Applying Advanced In Vitro Culturing Technology to Study the Human Gut Microbiota

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

Human Cartilage Tissue Fabrication Using Three-dimensional Inkjet Printing Technology

Bioprinting, which is based on thermal inkjet printing, is one of the most attractive enabling technologies in the field of tissue engineering and regenerative medicine. With digital control&#160;cells, scaffolds, and growth factors can be precisely deposited to the desired two-dimensional (2D) and three-dimensional (3D) locations rapidly. Therefore, this technology is an ideal approach to fabricate tissues mimicking their native anatomic structures. In order to engineer cartilage with native zonal organization, extracellular matrix composition (ECM), and mechanical properties, we developed a bioprinting platform using a commercial inkjet printer with simultaneous photopolymerization capable for 3D cartilage tissue engineering. Human chondrocytes suspended in poly(ethylene glycol) diacrylate (PEGDA) were printed for 3D neocartilage construction via layer-by-layer assembly. The printed cells were fixed at their original deposited positions, supported by the surrounding scaffold in simultaneous photopolymerization. The mechanical properties of the printed tissue were similar to the native cartilage. Compared to conventional tissue fabrication, which requires longer UV exposure, the viability of the printed cells with simultaneous photopolymerization was significantly higher. Printed neocartilage demonstrated excellent glycosaminoglycan (GAG) and collagen type II production, which was consistent with gene expression. Therefore, this platform is ideal for accurate cell distribution and arrangement for anatomic tissue engineering.

The methods described in this paper show how to convert a commercial inkjet printer into a bioprinter with simultaneous UV polymerization. The printer is capable of constructing 3D tissue structure with cells and biomaterials. The study demonstrated here constructed a 3D neocartilage.

Bioprinting, which is based on thermal inkjet printing, is one of the most attractive enabling technologies in the field of tissue engineering and ...

Culturing Mouse Cardiac Valves in the Miniature Tissue Culture System

Heart valve disease is a major burden in the Western world and no effective treatment is available. This is mainly due to a lack of knowledge of the molecular, cellular and mechanical mechanisms underlying the maintenance and/or loss of the valvular structure. 
Current models used to study valvular biology include in vitro cultures of valvular endothelial and interstitial cells. Although, in vitro culturing models provide both cellular and molecular mechanisms, the mechanisms involved in the 3D-organization of the valve remain unclear. While in vivo models have provided insight into the molecular mechanisms underlying valvular development, insight into adult valvular biology is still elusive. 
In order to be able to study the regulation of the valvular 3D-organization on tissue, cellular and molecular levels, we have developed the Miniature Tissue Culture System. In this ex vivo flow model the mitral or the aortic valve is cultured in its natural position in the heart. The natural configuration and composition of the leaflet are maintained allowing the most natural response of the valvular cells to stimuli. The valves remain viable and are responsive to changing environmental conditions. This MTCS may provide advantages on studying questions including but not limited to, how does the 3D organization affect valvular biology, what factors affect 3D organization of the valve, and which network of signaling pathways regulates the 3D organization of the valve.

Here, we present an ex vivo flow model in which murine cardiac valves can be cultured allowing the study of the biology of the valve.

Heart valve disease is a major burden in the Western world and no effective treatment is available. This is mainly due to a lack of knowledge of the ...

Eye Irritation Test (EIT) for Hazard Identification of Eye Irritating Chemicals using Reconstructed Human Cornea-like Epithelial (RhCE) Tissue Model

To comply with the Seventh Amendment to the EU Cosmetics Directive and EU REACH legislation, validated non-animal alternative methods for reliable and accurate assessment of ocular toxicity in man are needed. To address this need, we have developed an eye irritation test (EIT) which utilizes a three dimensional reconstructed human cornea-like epithelial (RhCE) tissue model that is based on normal human cells. The EIT is able to separate ocular&#160;irritants and corrosives (GHS Categories 1 and 2 combined) and those that do not require labeling (GHS No Category). The test utilizes two separate protocols, one designed for liquid chemicals and a second, similar protocol for solid test articles. The EIT prediction model uses a single exposure period (30 min for liquids, 6 hr for solids) and a single tissue viability cut-off (60.0% as determined by the MTT assay). Based on the results for 83 chemicals (44 liquids and 39 solids) EIT achieved 95.5/68.2/ and 81.8% sensitivity/specificity and accuracy (SS&#38;A) for liquids, 100.0/68.4/ and 84.6% SS&#38;A for solids, and 97.6/68.3/ and 83.1% for overall SS&#38;A. The EIT will contribute significantly to classifying the ocular irritation potential of a wide range of liquid and solid chemicals without the use of animals to meet regulatory testing requirements. The EpiOcular EIT method was implemented in 2015 into the OECD Test Guidelines as TG 492.

Eye Irritation Test EIT for Hazard Identification of Eye Irritating Chemicals using Reconstructed Human Cornea-like Epithelial RhCE Tissue Model

We have developed an eye irritation test which utilizes a three dimensional reconstructed human cornea-like epithelial (RhCE) tissue model. The test is able to discriminate between ocular irritant and corrosive materials (GHS Categories 1 and 2 combined) and those that do not require labeling (GHS No Category).

To comply with the Seventh Amendment to the EU Cosmetics Directive and EU REACH legislation, validated non-animal alternative methods for reliable and ...

TAPE: A Biodegradable Hemostatic Glue Inspired by a Ubiquitous Compound in Plants for Surgical Application

This video describes the simplest protocol for preparing biodegradable surgical glue that has an effective hemostatic ability and greater water-resistant adhesion strength than commercial tissue adhesives. Medical adhesives have attracted great attention as potential alternative tools to sutures and staples due to their convenience in usage with minimal invasiveness. Although there are several protocols for developing tissue adhesives including those commercially available such as fibrin glues and cyanoacrylate-based materials, mostly they require a series of chemical syntheses of organic molecules, or complicated protein-purification methods, in the case of bio-driven materials (i.e., fibrin glue). Also, the development of surgical glues exhibiting high adhesive properties while maintaining biodegradability is still a challenge due to difficulties in achieving good performance in the wet environment of the body. We illustrate a new method to prepare a medical glue, known as TAPE, by the weight-based separation of a water-immiscible supramolecular aggregate formed after a physical mixing of a plant-derived, wet-resistant adhesive molecule,&#160;Tannic Acid (TA), and a well-known biopolymer, Poly(Ethylene) glycol (PEG). With our approach, TAPE shows high adhesion strength, which is 2.5-fold more than commercial fibrin glue in the presence of water. Furthermore, TAPE is biodegradable in physiological conditions and can be used as a potent hemostatic glue against tissue bleeding. We expect the widespread use of TAPE in a variety of medical settings and drug delivery applications, such as polymers for muco-adhesion, drug depots, and others.

We describe the simplest protocol to prepare biodegradable medical glue that has an effective hemostatic ability. TAPE is a water-immiscible supramolecular aggregate prepared by mixing of tannic acid, a ubiquitous compound found in plants, and poly(ethylene) glycol, yielding a 2.5 times greater water-resistant adhesion compared with commercial fibrin glue.

This video describes the simplest protocol for preparing biodegradable surgical glue that has an effective hemostatic ability and greater water-resistant ...

Co-culture of Living Microbiome with Microengineered Human Intestinal Villi in a Gut-on-a-Chip Microfluidic Device

Here, we describe a protocol to perform long-term co-culture of multi-species human gut microbiome with microengineered intestinal villi in a human gut-on-a-chip microphysiological device. We recapitulate the intestinal lumen-capillary tissue interface in a microfluidic device, where physiological mechanical deformations and fluid shear flow are constantly applied to mimic peristalsis. In the lumen microchannel, human intestinal epithelial Caco-2 cells are cultured to form a &#39;germ-free&#39; villus epithelium and regenerate small intestinal villi. Pre-cultured microbial cells are inoculated into the lumen side to establish a host-microbe ecosystem. After microbial cells adhere to the apical surface of the villi, fluid flow and mechanical deformations are resumed to produce a steady-state microenvironment in which fresh culture medium is constantly supplied and unbound bacteria (as well as bacterial wastes) are continuously removed. After extended co-culture from days to weeks, multiple microcolonies are found to be randomly located between the villi, and both microbial and epithelial cells remain viable and functional for at least one week in culture. Our co-culture protocol can be adapted to provide a versatile platform for other host-microbiome ecosystems that can be found in various human organs, which may facilitate in vitro study of the role of human microbiome in orchestrating health and disease.

We describe an in vitro protocol to co-culture gut microbiome and intestinal villi for an extended period using a human gut-on-a-chip microphysiological system.

Here, we describe a protocol to perform long-term co-culture of multi-species human gut microbiome with microengineered intestinal villi in a human ...

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and dampen loads in the spine. The IVD consists of a proteoglycan-rich nucleus pulposus (NP) and a collagen-rich annulus fibrosis (AF) sandwiched by cartilaginous end-plates. Together with the adjacent vertebrae, the vertebrae-IVD structure forms a functional spine unit (FSU). These microstructures contain unique cell types as well as unique extracellular matrices. Whole organ culture of the FSU preserves the native extracellular matrix, cell differentiation phenotypes, and cellular-matrix interactions. Thus, organ culture techniques are particularly useful for investigating the complex biological mechanisms of the IVD. Here, we describe a high-throughput approach for culturing whole lumbar mouse FSUs that provides an ideal platform for studying disease mechanisms and therapies for the IVD. Furthermore, we describe several applications that utilize this organ culture method to conduct further studies including contrast-enhanced microCT imaging and three-dimensional high-resolution finite element modeling of the IVD.

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Whole organ culture of the intervertebral disc (IVD) preserves the native extracellular matrix, cell phenotypes, and cellular-matrix interactions. Here we describe an IVD culture system using mouse lumbar and caudal IVDs in their functional spinal units and several applications utilizing this system.

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and ...

Melt Electrospinning Writing of Three-dimensional Poly(&#949;-caprolactone) Scaffolds with Controllable Morphologies for Tissue Engineering Applications

This tutorial reflects on the fundamental principles and guidelines for electrospinning writing with polymer melts, an additive manufacturing technology with great potential for biomedical applications. The technique facilitates the direct deposition of biocompatible polymer fibers to fabricate well-ordered scaffolds in the sub-micron to micro scale range. The establishment of a stable, viscoelastic, polymer jet between a spinneret and a collector is achieved using an applied voltage and can be direct-written. A significant benefit of a typical porous scaffold is a high surface-to-volume ratio which provides increased effective adhesion sites for cell attachment and growth. Controlling the printing process by fine-tuning the system parameters enables high reproducibility in the quality of the printed scaffolds. It also provides a flexible manufacturing platform for users to tailor the morphological structures of the scaffolds to their specific requirements. For this purpose, we present a protocol to obtain different fiber diameters using melt electrospinning writing (MEW) with a guided amendment of the parameters, including flow rate, voltage and collection speed. Furthermore, we demonstrate how to optimize the jet, discuss often experienced technical challenges, explain troubleshooting techniques and showcase a wide range of printable scaffold architectures.

Melt Electrospinning Writing of Three-dimensional Poly(&amp;#949;-caprolactone) Scaffolds with Controllable Morphologies for Tissue Engineering Applications

This protocol serves as a comprehensive guideline to fabricate scaffolds via electrospinning with polymer melts in a direct writing mode. We systematically outline the process and define the appropriate parameter settings for achieving targeted scaffold architectures.

This tutorial reflects on the fundamental principles and guidelines for electrospinning writing with polymer melts, an additive manufacturing technology ...

Creating a Structurally Realistic Finite Element Geometric Model of a Cardiomyocyte to Study the Role of Cellular Architecture in Cardiomyocyte Systems Biology

With the advent of three-dimensional (3D) imaging technologies such as electron tomography, serial-block-face scanning electron microscopy and confocal microscopy, the scientific community has unprecedented access to large datasets at sub-micrometer resolution that characterize the architectural remodeling that accompanies changes in cardiomyocyte function in health and disease. However, these datasets have been under-utilized for&#160;investigating the role of cellular architecture remodeling in cardiomyocyte function. The purpose of this protocol is to outline how to create an accurate finite element model of a cardiomyocyte using high resolution electron microscopy and confocal microscopy images. A detailed and accurate model of cellular architecture has significant potential to provide new insights into cardiomyocyte biology, more than experiments alone can garner. The power of this method lies in its ability to computationally fuse information from two disparate imaging modalities of cardiomyocyte ultrastructure to develop one unified and detailed model of the cardiomyocyte. This protocol outlines steps to integrate electron tomography and confocal microscopy images of adult male Wistar (name for a specific breed of albino rat) rat cardiomyocytes to develop a half-sarcomere finite element model of the cardiomyocyte. The procedure generates a 3D finite element model that contains an accurate, high-resolution depiction (on the order of ~35 nm) of the distribution of mitochondria, myofibrils and ryanodine receptor clusters that release the necessary calcium for cardiomyocyte contraction from the sarcoplasmic reticular network (SR) into the myofibril and cytosolic compartment. The model generated here as an illustration does not incorporate details of the transverse-tubule architecture or the sarcoplasmic reticular network and is therefore a minimal model of the cardiomyocyte. Nevertheless, the model can already be applied in simulation-based investigations into the role of cell structure in calcium signaling and mitochondrial bioenergetics, which is illustrated and discussed using two case studies that are presented following the detailed protocol.

This protocol outlines a novel method to create a spatially detailed finite element model of the intracellular architecture of cardiomyocytes from electron microscopy and confocal microscopy images. The power of this spatially detailed model is demonstrated using case studies in calcium signaling and bioenergetics.

With the advent of three-dimensional (3D) imaging technologies such as electron tomography, serial-block-face scanning electron microscopy and confocal ...

Bioparticle Microarrays for Chemotactic and Molecular Analysis of Human Neutrophil Swarming in vitro

Neutrophil swarming is a cooperative process by which neutrophils seal off a site of infection and promote tissue reorganization. Swarming has classically been studied in vivo in animal models showing characteristic patterns of cell migration. However, in vivo models have several limitations, including intercellular mediators that are difficult to access and analyze, as well as the inability to directly analyze human neutrophils. Because of these limitations, there is a need for an in vitro platform that studies swarming with human neutrophils and provides easy access to the molecular signals generated during swarming. Here, a multistep microstamping process is used to generate a bioparticle microarray that stimulates swarming by mimicking an in vivo infection. The bioparticle microarray induces neutrophils to swarm in a controlled and stable manner. On the microarray, neutrophils increase in speed and form stable swarms around bioparticle clusters. Additionally, supernatant generated by the neutrophils was analyzed and 16 proteins were discovered to have been differentially expressed over the course of swarming. This in vitro swarming platform facilitates direct analysis of neutrophil migration and protein release in a reproducible, spatially controlled manner.

This protocol generates bioparticle microarrays that provide spatially controlled neutrophil swarming. It provides easy access to the mediators that neutrophils release during migration and allows for quantitative imaging analysis.

Neutrophil swarming is a cooperative process by which neutrophils seal off a site of infection and promote tissue reorganization. Swarming has classically ...