Thanks to the Umberto Veronesi Foundation for a fellowship supporting researcher work.

1. Preparation of the basic 3D reconstructed intestinal mucosa model
NOTE: The entire procedure must be carried out in a sterile laminar flow hood. All steps in the procedure involving the use of the cell incubator signify that cultures are incubated at 37 &#176;C in a humidified atmosphere containing 5% CO2 (unless stated otherwise in the protocol).
<ol>
	<li>Prior preparation of the cell lines used in the intestinal model system
	<ol>
		<li>Seed L929 mouse fibroblast cells at a concentration of 5 x 105 in 5 mL of Dulbecco&#39;s Modified Eagle Medium (DMEM) containing 2 mM L-glutamine, 10% Fetal Bovine Serum (FBS), and 1% Penicillin-Streptomycin (Pen-Strep) in a&#160;F25 flask and culture it in an incubator 4 days before the construction of the intestinal models. After 48 h, remove the medium using a pipette. Then add fresh medium (5 mL), and incubate the cells further for 48 h. 
		NOTE: The cells must be 80% confluent before being used.</li>
		<li>Seed Caco-2 cells (concentration of 2 x 106) in 10 mL of DMEM medium (with 10% FBS and 1% Pen-Strep) in a&#160;F75 cell flask and culture it in an incubator 4 days before the construction of the intestinal mucosa model. After 48 h, change the medium as described in step 1.1.1 and further incubate for 48 h to a confluency of 80%. 
		NOTE: The cells must be in an active proliferating phase: not too sparse nor too confluent. A 50-60% confluence is recommended. The cells must not be sown the day before making the model because this could slow down the proliferative capacity of the cells, which would then not take root perfectly in the reconstructed 3D model.</li>
		<li>Add U937 cells that grow in suspension (concentration of 1 x 106) to 10 mL of Roswell Park Memorial Institute (RPMI) medium (containing 2 mM L-glutamine, 1% sodium pyruvate, 10% FBS, and 1% Pen-Strep) in a F75 cell flask 2 days prior to the model set-up, and place in an incubator for 48 h.</li>
	</ol>
	</li>
	<li>Preparation of the Transwell co-culture plate inserts
	<ol>
		<li>Select a 24-well plate containing inserts with 0.4 &#181;m filters. 
		NOTE: The 0.4 &#181;m filter is a standard choice in drug transport studies. The 3 &#181;m and 8 &#181;m filter sizes are not recommended to prevent any possible losses of the collagen-embedded cells.</li>
		<li>Using a pipette, hydrate the Transwell filters (henceforth referred to as membrane inserts) with 500 &#181;L of Hanks&#39; Balanced Salt Solution (HBSS) below and above the filter insert.</li>
		<li>Close the multi-well plate and place it in an incubator for a minimum of 2 h. 
		NOTE: The membrane inserts can remain in the HBSS for up to 24 h. This operation can be performed the day before constructing the model. It is important that the membrane inserts are completely hydrated and that the filter does not dry out, as this could make it harder for the sample to adhere properly.</li>
		<li>Remove the plates from the incubator after 2 h (or 24 h). Carefully aspirate the HBSS from above and below the membrane inserts using a pipette and leave it to dry for 10 min.</li>
	</ol>
	</li>
	<li>Preparation of the cell-free collagen lamina propria of the basic 3D intestinal model system (DAY 1)
	<ol>
		<li>Prepare a cell-free collagen solution in a 50 mL sterile tube on ice containing the following components in DMEM: 10% Fetal Bovine Serum (FBS), 200 mM L-glutamine, 1% sodium bicarbonate and 1.35 mg/mL Type 1 rat tail collagen. 
		NOTE: All these components must be kept on ice and added to the DMEM with cooled pipette tips. The Type 1 rat tail collagen must be added last as this polymerizes with increasing temperature and pH. The amount of cell-free collagen solution prepared will depend on the number of intestinal equivalents required.</li>
		<li>Add 250 &#181;L of the solution to each plate insert (above the membrane filter) for the number of intestinal equivalents selected and place the lid over the multi-well plate. Allow the collagen solution to polymerize at room temperature (RT) under the laminar flow hood. 
		NOTE: Apart from the transition to a more solid phase, polymerization is also evident from a change in color from yellow to pink. Polymerization is usually complete within 10-15 min.</li>
	</ol>
	</li>
	<li>Preparation, cell counting, and addition of fibroblast (L929) and monocyte (U937) cells to the model system (DAY 1)
	<ol>
		<li>Remove the L929 cell culture (step 1.1.1) from the incubator. Using a vacuum pump, aspirate the medium, replace it with 5 mL of sterile phosphate buffer saline (PBS; without Ca and Mg), and rinse the cells.</li>
		<li>Aspirate the PBS using a vacuum pump. Add 2 mL of a pre-prepared trypsin-EDTA solution (0.05% trypsin and 0.02% EDTA in PBS) and place in an incubator for 3-5 min.</li>
		<li>Use an inverted microscope (for example, an Eclipse Ts2, Nikon) to establish whether the cells are detaching from the adhesion surface. If this is occurring, immediately add 2 mL of DMEM (containing 10% FBS) to block the trypsin reaction and rinse the cells.</li>
		<li>Transfer the cell solution to a sterile 15 mL tube and centrifuge at 645 x g for 5 min. Using a vacuum pump, aspirate the supernatant. 
		NOTE: Care is needed not to disturb the pellet. The effect of DMEM was tested on the L929 and U937 cells and was not shown to have adverse effects on growth.</li>
		<li>Add 1 mL of DMEM to the pellet and suspend the cells. 
		NOTE: The cells must be homogeneously suspended in the solution.</li>
		<li>Remove 20 &#181;L of the cells in DMEM and add 20 &#181;L of Trypan blue solution. Remove 20 &#181;L of the mix and evaluate cell density microscopically using a cell counting chamber.</li>
		<li>Remove the U937 cell culture (step 1.1.3) from the incubator. Centrifuge the cell solution at 645 x g for 5 min. Using a vacuum pump, aspirate the supernatant to avoid disturbing the pellet.</li>
		<li>Suspend the cells in 1 mL of RPMI. As with the L929 cells, make sure the cells are homogeneously suspended in the solution.</li>
		<li>Similarly, establish the cell density as reported in step 1.4.6.</li>
		<li>Prepare a collagen solution as described in step 1.3.1. 
		NOTE: The amount of solution to be made must take into consideration 450 &#181;L for each insert (or model).</li>
		<li>Prepare a solution of DMEM to contain a count of 50,000 L929 cells and 15,000 U937 cells, respectively, in a volume of 50 &#181;L for each intestinal equivalent model to be constructed. 
		NOTE: The number of cells is a critical factor. Too many of both cell types would result in a lamina propria excessively full of cells that would not be adequately organized. An inferior number of fibroblasts (which generate collagen) would result in a less compact lamina propria, whereas too few monocytes would impede the immune response to stimuli. Provided the correct count of cells is present within each 50 &#181;L aliquot - a total volume of 600 &#181;L can be prepared for the 12 filter inserts present in each 24 well-plate.</li>
		<li>Add each 50 &#181;L aliquot containing the cells to 450 &#181;L of collagen solution in step 1.4.10. Mix thoroughly.</li>
		<li>Overlay the pre-coated cell-free collagen lamina propria with 500 &#181;L of the collagen-containing cell solution for each model. 
		NOTE: It is important to rapidly add the volumes to each insert, and as such, it is advisable to limit the number of reconstructed intestinal mucosa models to 12 or fewer at a time.</li>
		<li>Close the plate and place it in an incubator for 2 h to allow the solution to set.</li>
	</ol>
	</li>
	<li>Preparation, cell counting, and addition of epithelial Caco-2 cells to the intestinal model (DAY 1)
	<ol>
		<li>Remove the Caco-2 cell culture (step 1.1.2) from the incubator. Repeat the procedures from step 1.4.1 using 10 mL of PBS for preliminary rinsing. Then add 5 mL of a pre-prepared trypsin-EDTA solution (0.05% trypsin and 0.02% EDTA in Ca- and Mg-free PBS) and place in an incubator for 5-8 min.</li>
		<li>Repeat steps 1.4.3 (but using 5 mL of DMEM to block the trypsin reaction) through to 1.4.6 to count the cells using the cell counting chamber.</li>
		<li>Prepare a solution of DMEM to contain a count of 150,000 Caco-2 cells in 50 &#181;L. 
		NOTE: The number of cells used is a critical point. Exceeding 150,000 cells could result in a compact disorganized epithelial layer, whereas with too little (less than 100,000), the cells struggle to grow and do not adequately cover the basement membrane creating a discontinuous intestinal epithelial layer. Make sure the cells are effectively suspended. The tip of a 200 &#181;L pipette can be used to ensure homogenous distribution. Given that 12 models can be constructed at any given time, a 600 &#181;L cell solution can be prepared.</li>
		<li>After the 2 h required in step 1.4.14, add 50 &#181;L of Caco-2 cells suspended in DMEM to the middle of each basement membrane. Close the lid of the multi-well plate.</li>
		<li>Incubate under the sterile laminar flow hood for 10 min. Then transfer to the incubator for 30 min.</li>
		<li>Prepare a DMEM solution containing 10% FBS and 1% Pen/Strep. 
		NOTE: Prepare a sufficient solution to use a 1 mL volume per model.</li>
		<li>Add 500 &#181;L of the medium above the reconstructed model and 500 &#181;L below the filter. 
		NOTE: Care is required when adding the medium above the filter to avoid detaching or stressing the cells.</li>
		<li>Close the multi-well plate system and place it in the incubator.</li>
	</ol>
	</li>
	<li>Model preparation/formation and use (DAY 2 to DAY 6)
	<ol>
		<li>DAY 2: Carefully remove the solution from both above the reconstructed model and below the filter using a pipette. Replace with fresh 500 &#181;L of fresh DMEM (10% FBS and 1% Pen/Strep) above and below the filter, respectively.</li>
		<li>DAY 3: Repeat as above in step 1.6.1.</li>
		<li>DAY 4: Repeat as above in step 1.6.1. 
		NOTE: This intestinal model is a static cellular model; therefore, to favor the release of molecules and the growth and stimulation of cells, it is very important that the medium is changed every day.</li>
		<li>DAY 5: At this point, the model is fully formed/developed. Use these models for further studies. 
		NOTE: The best time to use the model is at 5 days. Although the cells can be maintained in an incubator for longer periods, the more time that passes, the greater the probability that the epithelial cells may grow in an uncontrolled manner, resulting in an unorganized and compact layer that is difficult to use.</li>
		<li>At 5 days, incubate the models with either toxic (gluten or lipopolysaccharide [LPS]) or beneficial components (polyphenols) of interest. Add these to the upper portion of the reconstructed model suspended in the DMEM medium. 
		NOTE: The suitable concentration of each component of interest must be calculated and suspended in a DMEM medium. Non-treated controls containing only DMEM medium must be set up for comparison to the experimental models.</li>
		<li>Incubate the control and experimental models for 24 h in an incubator.</li>
		<li>DAY 6: Remove the medium above and below the filter with a pipette. 
		&#8203;NOTE: The medium can be stored for subsequent enzyme-linked immunosorbent assay (ELISA)-type tests to measure the release of inflammatory cytokines. For this purpose, the medium must be added to sterile vials and stored at -20 &#176; C for further analysis.</li>
	</ol>
	</li>
</ol>
2. Paraffin embedding of the reconstructed intestinal mucosa models
NOTE: The entire procedure must be carried out under a chemical fume hood. Each step and respective time allocation must be strictly adhered to. For this reason, it is important to have all reagents prepared ahead of time.
<ol>
	<li>Set the paraffin machine to 58 &#176;C so that&#160;it is ready for use.</li>
	<li>Transfer the membrane inserts to clean wells using pliers in a sterile 24-well plate.</li>
	<li>Add 500 &#181;L of 37% buffered formalin in PBS above the filter and 1 mL under the filter. Close the lid and leave it under the chemical fume hood for 2 h at RT. 
	NOTE: As an alternative, 4% buffered formalin can be used at RT for 1 h.</li>
	<li>Remove the formalin and add HBSS solution both above and below the filter. Then remove the HBSS.</li>
	<li>Detach the lamina propria from the membrane insert. 
	NOTE: The cells of the intestinal mucosa usually detach very easily from the membrane insert as they are not attached to the latter. Additionally, the two sample compartments (apical and basal) remain attached, retaining the 3D structure. If, for some reason, they are not readily detached, it will be important to use a sterile disposable scalpel blade to remove the intestinal mucosa from the membrane insert. The membrane insert can be cut, thus detaching it from the plastic. Be careful never to touch the sample with the scalpel. The motive for removing the membrane insert from the collagen-embedded cells is that this filter may become detached in the subsequent fixation or paraffin embedding phases, rendering comparisons between intestinal mucosa models disproportional. Moreover, the membrane inserts within the embedded section have a different consistency which can interfere with sectioning.</li>
	<li>Place the 100 mL beakers under the chemical fume hood, each correctly labeled to include one reconstructed intestinal mucosa model system under investigation. Add 25 mL of 35% ETOH to each beaker and then add the reconstructed intestinal mucosa. Incubate for 10 min.</li>
	<li>Replace the 35% ethanol with 25 mL of 50% ethanol and incubate for 10 min.</li>
	<li>Replace the 50% ethanol with 25 mL of 70% ethanol and incubate for 10 min.</li>
	<li>Replace the 70% ethanol with 25 mL of 80% ethanol and incubate for 10 min.</li>
	<li>Replace the 80% ethanol with 25 mL of 95% ethanol and incubate for 10 min.</li>
	<li>Replace the 95% ethanol with 25 mL of 100% ethanol and incubate for 10 min.</li>
	<li>Replace the 100% ethanol with another 25 mL change of 100% ethanol and incubate for 10 min.</li>
	<li>Place 100 mL beakers under the fume hood, each correctly labeled to include one model under investigation. Add 50 mL of xylene or a histological clearing agent for 10-20 min. 
	NOTE: Histo-Clear (histological clearing agent) is recommended as the agent enhances the clarity and vibrancy of acidophilic stains. The clearing time can vary from one reconstructed intestinal mucosa sample to another and must be checked for transparency in appearance every 2-3 min.</li>
	<li>As soon as the samples are transparent, place them in a metal tissue cassette holder which is then immersed in liquid paraffin inside the heated machine for 45 min.</li>
	<li>Change the paraffin and leave the samples inside the heated machine for a further 45 min.</li>
	<li>Remove the tissue cassette holder and place on ice to cool. When the cooled sample blocks detach from the metal holder, these can be further cooled at room temperature.</li>
	<li>Store the blocks at RT. 
	NOTE: If the objective is to prepare sections immediately, then the blocks can be placed at 4 &#176; C so that they are well-cooled when used.</li>
	<li>Cut 4 &#181;m sections using a microtome.</li>
	<li>Place the cut sections on slides and dry them in an oven at 37 &#176; C for 24 h.</li>
	<li>The slides are ready for use. Store them at RT until performing either H&#38;E histological staining or immune-histochemical reaction staining (antigen-antibody type).
	<ol>
		<li>For immune-histochemical staining, select the antibodies of interest (either for the study of TJ proteins&#160;such as occludin or for monocyte activation, migration, and differentiation) and detect expression (via staining) using commercial kits.</li>
		<li>Similarly, measure mucus using the Alcian blue and periodic acid-Schiff (PAS) staining kit.</li>
		<li>Quantify the stained TJ proteins of interest, such as occludin, by calculating the percentage of positive pixels on micrographs taken from the microscope.</li>
		<li>Analyze pictures of the cells using image analysis software (e.g., ImageJ2 software [Wayne Rasband, version 2.9.0/1.53t]).
		<ol>
			<li>To perform the analysis of the pixels, process the digital images to 300 pixels/inch and convert them to 8 bits. Then, process the binary images by Color Deconvolution plugin to analyze the staining of the protein of interest, in this case, the permanent red staining of occludin.</li>
			<li>Save the selected picture as a tiff and subject it to a &#34;clean-up&#34; procedure to eliminate artifacts with a graphics editor (e.g., Adobe PhotoshopCC [version 20.0.4]).</li>
			<li>Thereafter, measure all fields of interest with the application Analyze Particle of ImageJ2, and report the data as the number of pixels. 
			NOTE: Each experiment should be performed in triplicate with three internal replicate fields analyzed for each replicate. For quantifying mucus, the same principle of measuring the pixels can be applied to the images taken of the purple-magenta-stained neutral mucins and bright blue-stained acidic mucus, respectively.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

Construction of a 3D Intestinal Mucosa Model with Paraffin Embedding

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The authors declare no conflict of interest.

The basic reconstructed intestinal mucosa model system presented here (Figure 6) combines physiological complexity (more physiologically relevant 3D cell cultures containing a Caco-2 monolayer with an ECM-rich lamina propria support containing fibroblasts and monocytes) with experimental simplicity (using commercial human cell lines to produce standardized and easily repeatable system)13. As such, this model system is deemed a suitable alternative to murine models aim...

The intestinal epithelial barrier, a one-cell-thick internal lining containing different types of epithelial cells, constitutes the first physical defensive barrier or interface between the outside and the internal milieu of the body1,2. Columnar-type enterocytes constitute the most abundant type of epithelial cells. These are&#160;responsible for maintaining epithelial barrier integrity through interactions between several barrier components, including tight junctions (TJs), playing a significant role in barrier tightening1,3. The TJ structure consists of intracellular plaque proteins, such as zonula occludens (ZO) and cingulin, cooperating with transmembrane proteins, including occludins, claudins, and junctional adhesion molecules (JAMs) that form zipper-like structure tightly linking the neighboring cells3,4. The transmembrane proteins regulate the passive paracellular diffusion of small compounds and exclude toxic large molecules.
Potentially toxic food compounds and food contaminants stimulate inflammatory cytokine production that disrupts the epithelial permeability, activating immune cells and causing chronic intestinal tissue inflammation5,6,7. In contrast, various anti-oxidant and anti-inflammatory phytochemicals have been reported to reduce inflammatory cytokine expression and enhance intestinal TJ barrier integrity through the restoration of TJ protein expression and assembly4,6,8. Hence, the regulation of epithelial barrier integrity by both beneficial and harmful compounds has seen an increase in the use of both in vivo and in vitro models aimed at mimicking the intestinal barrier for pharmaceutical screening and toxicity studies. This is particularly relevant given the increasing interest in understanding the pathophysiology of intestinal bowel diseases (IBD), necrotizing enterocolitis, and cancer, which can be simulated in experimental models8,9,10.
There has been a demand for the development of cell-based in vitro models in order to achieve the objective of the &#8220;3Rs&#8221; in animal testing. These include replacement alternatives to the use of animals, reduction in the number of animals used, and refinement in adopting methods that alleviate distress11,12,13. Moreover, the underlying molecular, cellular, and physiological mechanisms between human and murine models (rodents being the most widely used species) are distinctive, leading to controversy regarding the efficacy of the murine models as predictors in human responses12,13. Numerous advantages of in vitro human cell-line models include target-restricted experimentation, direct observation, and continuous analysis13.
Single-cell-type monolayers in two-dimensional (2D) cultures have served as powerful models. However, these cannot precisely reproduce the physiological complexity of human tissues8,13,14. As a result, 3D&#160;culture systems are being developed with ever-increasing improvements to recapitulate the physiological complexity of both healthy and diseased intestinal tissues as next-generation risk assessment toolboxes13,14. These models include 3D Transwell scaffolds with diverse cell lines, organoid cultures, and microfluidic devices (intestine-on-chip) using both cell lines and organoids (derived from both healthy and diseased tissues)8,13,14.&#160;
The 3D &#8220;healthy tissue&#8221; intestinal equivalent protocol presented in the present study was based on striking a balance between physiological complexity and experimental simplicity13. The model is representative of a 3D Transwell scaffold, comprised of three cell lines (enterocytes [the gold-standard colon adenocarcinoma Caco-2 line] with a supporting immune component [U937 monocytes and L929 fibroblasts]), constituting a standardized and easily repeatable system applicable for the preliminary screening of dietary molecules of interest on intestinal epithelial barrier integrity and immune response. The protocol includes paraffin embedding for light microscopic evaluation of epithelial barrier integrity using fixed intestinal equivalents. The advantage of the present approach is that numerous sections of the embedded tissues can be made to stain for multiple parameters from a single experiment.

basic three-dimensional (3d) intestinal model system with an immune component

There has been an increase in the use of in vivo and in vitro intestinal models to study the pathophysiology of inflammatory intestinal diseases, for the pharmacological screening of potentially beneficial substances, and for toxicity studies on potentially harmful food components. Of relevance, there is a current demand for the development of cell-based in vitro models to substitute animal models. Here, a protocol for a basic, &#8220;healthy tissue&#8221; three-dimensional (3D) intestinal equivalent model using cell lines is presented with the dual benefit of providing both experimental simplicity (standardized and easily repeatable system) and physiological complexity (Caco-2 enterocytes with a supporting immune component of U937 monocytes and L929 fibroblasts). The protocol also includes paraffin embedding for light microscopic evaluation of fixed intestinal equivalents, thereby providing the advantage of analyzing multiple visual parameters from a single experiment. Hematoxylin and eosin (H&#38;E) stained sections showing the Caco-2 columnar cells forming a tight and regular monolayer in control treatments are used to verify the efficacy of the model as an experimental system. Using gluten as a pro-inflammatory food component, parameters analyzed from sections include reduced monolayer thickness, as well as disruption and detachment from the underlying matrix (H&#38;E), decreased tight junction protein expression as shown from occludin staining (quantifiable statistically), and immune-activation of migrating U937 cells as evidenced from the cluster of differentiation 14 (CD14) staining and CD11b-related differentiation into macrophages. As shown by using lipopolysaccharide to simulate intestinal inflammation, additional parameters that can be measured are increased mucus staining and cytokine expression (such as midkine) that can be extracted from the medium prior to fixation. The basic three-dimensional (3D) intestinal mucosa model and fixed sections can be recommended for inflammatory status and barrier integrity studies with the possibility of analyzing multiple visual quantifiable parameters.

The first important aspect is to determine the acceptability of the basic 3D intestinal equivalent mucosa for experimental purposes. This is performed with the most widely used stain in histology and histopathology laboratories, namely hematoxylin (stains nuclear material deep blue-purple color) and eosin (stains cytoplasmic material varying shades of pink). The H&#38;E staining is first performed on an untreated control, which is cultured under the same conditions and timeframe as the experimental treatments. From visua...

Watch this Scientific Journal Video about Investigating the Effects of Compounds on Intestinal Tissue Using 3D Human Cell Line Models at JoVE.com

Author Spotlight: Investigating the Effects of Compounds on Intestinal Tissue Using 3D Human Cell Line Models 

Here we describe constructing a basic three-dimensional (3D) intestinal cell line model system and a paraffin embedding protocol for light microscopic evaluation of fixed intestinal equivalents. Staining of selected proteins permits the analysis of multiple visual parameters from a single experiment for potential use in preclinical drug screening studies.

Basic Three-Dimensional (3D) Intestinal Model System with an Immune Component

There has been an increase in the use of in vivo and in vitro intestinal models to study the pathophysiology of inflammatory intestinal diseases, for the ...

basic-three-dimensional-3d-intestinal-model-system-with-an-immune

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Biology

1.0K Views.  Alma Mater Studiorum-University of Bologna. Here we describe constructing a basic three-dimensional (3D) intestinal cell line model system and a paraffin embedding protocol for light microscopic evaluation of fixed intestinal equivalents. Staining of selected proteins permits the analysis of multiple visual parameters from a single experiment for potential use in preclinical drug screening studies.

Video: Author Spotlight: Investigating the Effects of Compounds on Intestinal Tissue Using 3D Human Cell Line Models 

Microfabrication of Chip-sized Scaffolds for Three-dimensional Cell cultivation

Using microfabrication technologies is a prerequisite to create scaffolds of reproducible geometry and constant quality for three-dimensional cell cultivation. These technologies offer a wide spectrum of advantages not only for manufacturing but also for different applications. The size and shape of formed cell clusters can be influenced by the exact and reproducible architecture of the microfabricated scaffold and, therefore, the diffusion path length of nutrients and gases can be controlled.1 This is unquestionably a useful tool to prevent apoptosis and necrosis of cells due to an insufficient nutrient and gas supply or removal of cellular metabolites.
Our polymer chip, called CellChip, has the outer dimensions of 2 x 2 cm with a central microstructured area. This area is subdivided into an array of up to 1156 microcontainers with a typical dimension of 300 m edge length for the cubic design (cp- or cf-chip) or of 300 m diameter and depth for the round design (r-chip).2
So far, hot embossing or micro injection moulding (in combination with subsequent laborious machining of the parts) was used for the fabrication of the microstructured chips. Basically, micro injection moulding is one of the only polymer based replication techniques that, up to now, is capable for mass production of polymer microstructures.3 However, both techniques have certain unwanted limitations due to the processing of a viscous polymer melt with the generation of very thin walls or integrated through holes. In case of the CellChip, thin bottom layers are necessary to perforate the polymer and provide small pores of defined size to supply cells with culture medium e.g. by microfluidic perfusion of the containers.
In order to overcome these limitations and to reduce the manufacturing costs we have developed a new microtechnical approach on the basis of a down-scaled thermoforming process. For the manufacturing of highly porous and thin walled polymer chips, we use a combination of heavy ion irradiation, microthermoforming and track etching. In this so called "SMART" process (Substrate Modification And Replication by Thermoforming) thin polymer films are irradiated with energetic heavy projectiles of several hundred MeV introducing so-called "latent tracks" Subsequently, the film in a rubber elastic state is formed into three dimensional parts without modifying or annealing the tracks. After the forming process, selective chemical etching finally converts the tracks into cylindrical pores of adjustable diameter.

We present two processes for the microfabrication of porous polymer chips for three-dimensional cell cultivation. The first one is hot embossing combined with a solvent vapour welding process. The second one uses a recently developed microthermoforming process combined with ion track technology leading to a significant simplification of manufacture.

Using microfabrication technologies is a prerequisite to create scaffolds of reproducible geometry and constant quality for three-dimensional cell ...

Culturing and Applications of Rotating Wall Vessel Bioreactor Derived 3D Epithelial Cell Models

Cells and tissues in the body experience environmental conditions that influence their architecture, intercellular communications, and overall functions. For in vitro cell culture models to accurately mimic the tissue of interest, the growth environment of the culture is a critical aspect to consider. Commonly used conventional cell culture systems propagate epithelial cells on flat two-dimensional (2-D) impermeable surfaces. Although much has been learned from conventional cell culture systems, many findings are not reproducible in human clinical trials or tissue explants, potentially as a result of the lack of a physiologically relevant microenvironment.
Here, we describe a culture system that overcomes many of the culture condition boundaries of 2-D cell cultures, by using the innovative rotating wall vessel (RWV) bioreactor technology. We and others have shown that organotypic RWV-derived models can recapitulate structure, function, and authentic human responses to external stimuli similarly to human explant tissues 1-6. The RWV bioreactor is a suspension culture system that allows for the growth of epithelial cells under low physiological fluid shear conditions. The bioreactors come in two different formats, a high-aspect rotating vessel (HARV) or a slow-turning lateral vessel (STLV), in which they differ by their aeration source. Epithelial cells are added to the bioreactor of choice in combination with porous, collagen-coated microcarrier beads (Figure 1A). The cells utilize the beads as a growth scaffold during the constant free fall in the bioreactor (Figure 1B). The microenvironment provided by the bioreactor allows the cells to form three-dimensional (3-D) aggregates displaying in vivo-like characteristics often not observed under standard 2-D culture conditions (Figure 1D). These characteristics include tight junctions, mucus production, apical/basal orientation, in vivo protein localization, and additional epithelial cell-type specific properties.
The progression from a monolayer of epithelial cells to a fully differentiated 3-D aggregate varies based on cell type1, 7-13. Periodic sampling from the bioreactor allows for monitoring of epithelial aggregate formation, cellular differentiation markers and viability (Figure 1D). Once cellular differentiation and aggregate formation is established, the cells are harvested from the bioreactor, and similar assays performed on 2-D cells can be applied to the 3-D aggregates with a few considerations (Figure 1E-G). In this work, we describe detailed steps of how to culture 3-D epithelial cell aggregates in the RWV bioreactor system and a variety of potential assays and analyses that can be executed with the 3-D aggregates. These analyses include, but are not limited to, structural/morphological analysis (confocal, scanning and transmission electron microscopy), cytokine/chemokine secretion and cell signaling (cytometric bead array and Western blot analysis), gene expression analysis (real-time PCR), toxicological/drug analysis and host-pathogen interactions. The utilization of these assays set the foundation for more in-depth and expansive studies such as metabolomics, transcriptomics, proteomics and other array-based applications. Our goal is to present a non-conventional means of culturing human epithelial cells to produce organotypic 3-D models that recapitulate the human in vivo tissue, in a facile and robust system to be used by researchers with diverse scientific interests.

A rotating cell culture system that allows epithelial cells to grow under physiological conditions resulting in 3-D cellular aggregate formation is described. The aggregates generated display in vivo-like characteristics not observed in conventional culture models and serve as a more accurate organotypic model system for a multitude of scientific investigations.

Cells and tissues in the body experience environmental conditions that influence their architecture, intercellular communications, and overall functions. ...

An Analytical Tool that Quantifies Cellular Morphology Changes from Three-dimensional Fluorescence Images

The most common software analysis tools available for measuring fluorescence images are for two-dimensional (2D) data that rely on manual settings for inclusion and exclusion of data points, and computer-aided pattern recognition to support the interpretation and findings of the analysis. It has become increasingly important to be able to measure fluorescence images constructed from three-dimensional (3D) datasets in order to be able to capture the complexity of cellular dynamics and understand the basis of cellular plasticity within biological systems. Sophisticated microscopy instruments have permitted the visualization of 3D fluorescence images through the acquisition of multispectral fluorescence images and powerful analytical software that reconstructs the images from confocal stacks that then provide a 3D representation of the collected 2D images. Advanced design-based stereology methods have progressed from the approximation and assumptions of the original model-based stereology1 even in complex tissue sections2. Despite these scientific advances in microscopy, a need remains for an automated analytic method that fully exploits the intrinsic 3D data to allow for the analysis and quantification of the complex changes in cell morphology, protein localization and receptor trafficking. 
Current techniques available to quantify fluorescence images include Meta-Morph (Molecular Devices, Sunnyvale, CA) and Image J (NIH) which provide manual analysis. Imaris (Andor Technology, Belfast, Northern Ireland) software provides the feature MeasurementPro, which allows the manual creation of measurement points that can be placed in a volume image or drawn on a series of 2D slices to create a 3D object. This method is useful for single-click point measurements to measure a line distance between two objects or to create a polygon that encloses a region of interest, but it is difficult to apply to complex cellular network structures. Filament Tracer (Andor) allows automatic detection of the 3D neuronal filament-like however, this module has been developed to measure defined structures such as neurons, which are comprised of dendrites, axons and spines (tree-like structure). This module has been ingeniously utilized to make morphological measurements to non-neuronal cells3, however, the output data provide information of an extended cellular network by using a software that depends on a defined cell shape rather than being an amorphous-shaped cellular model. To overcome the issue of analyzing amorphous-shaped cells and making the software more suitable to a biological application, Imaris developed Imaris Cell. This was a scientific project with the Eidgen&ouml;ssische Technische Hochschule, which has been developed to calculate the relationship between cells and organelles. While the software enables the detection of biological constraints, by forcing one nucleus per cell and using cell membranes to segment cells, it cannot be utilized to analyze fluorescence data that are not continuous because ideally it builds cell surface without void spaces. To our knowledge, at present no user-modifiable automated approach that provides morphometric information from 3D fluorescence images has been developed that achieves cellular spatial information of an undefined shape (Figure 1). 
We have developed an analytical platform using the Imaris core software module and Imaris XT interfaced to MATLAB (Mat Works, Inc.). These tools allow the 3D measurement of cells without a pre-defined shape and with inconsistent fluorescence network components. Furthermore, this method will allow researchers who have extended expertise in biological systems, but not familiarity to computer applications, to perform quantification of morphological changes in cell dynamics.

We developed a software platform that utilizes Imaris Neuroscience, ImarisXT and MATLAB to measure the changes in morphology of an undefined shape taken from three-dimensional confocal fluorescence of single cells. This novel approach can be used to quantify changes in cell shape following receptor activation and therefore represents a possible additional tool for drug discovery.

The most common software analysis tools available for measuring fluorescence images are for two-dimensional (2D) data that rely on manual settings for ...

Using an Automated 3D-tracking System to Record Individual and Shoals of Adult Zebrafish

Like many aquatic animals, zebrafish (Danio rerio) moves in a 3D space. It is thus preferable to use a 3D recording system to study its behavior. The presented automatic video tracking system accomplishes this by using a mirror system and a calibration procedure that corrects for the considerable error introduced by the transition of light from water to air. With this system it is possible to record both single and groups of adult zebrafish. Before use, the system has to be calibrated. The system consists of three modules: Recording, Path Reconstruction, and Data Processing. The step-by-step protocols for calibration and using the three modules are presented. Depending on the experimental setup, the system can be used for testing neophobia, white aversion, social cohesion, motor impairments, novel object exploration etc. It is especially promising as a first-step tool to study the effects of drugs or mutations on basic behavioral patterns. The system provides information about vertical and horizontal distribution of the zebrafish, about the xyz-components of kinematic parameters (such as locomotion, velocity, acceleration, and turning angle) and it provides the data necessary to calculate parameters for social cohesions when testing shoals.

The use of a 3D automatic video system that can track individual and groups of zebrafish is described. As application example we explore the effects of the NMDA-receptor antagonist MK-801 on shoals of zebrafish.

Like many aquatic animals, zebrafish (Danio rerio) moves in a 3D space. It is thus preferable to use a 3D recording system to study its behavior. The ...

FRET Imaging in Three-dimensional Hydrogels

Imaging of F&#246;rster resonance energy transfer (FRET)&#160;is a powerful tool for examining cell biology in real-time. Studies utilizing FRET commonly employ two-dimensional (2D) culture, which does not mimic the three-dimensional (3D) cellular microenvironment. A method to perform quenched emission FRET imaging using conventional widefield epifluorescence microscopy of cells within a 3D hydrogel environment is presented. Here an analysis method for ratiometric FRET probes that yields linear ratios over the probe activation range is described. Measurement of intracellular cyclic adenosine monophosphate (cAMP) levels is demonstrated in chondrocytes under forskolin stimulation using a probe for EPAC1 activation (ICUE1) and the ability to detect differences in cAMP signaling dependent on hydrogel material type, herein a photocrosslinking hydrogel (PC-gel, polyethylene glycol dimethacrylate) and a thermoresponsive hydrogel (TR-gel). Compared with 2D FRET methods, this method requires little additional work. Laboratories already utilizing FRET imaging in 2D can easily adopt this method to perform cellular studies in a 3D microenvironment. It can further be applied to high throughput drug screening in engineered 3D microtissues. Additionally, it is compatible with other forms of FRET imaging, such as anisotropy measurement and fluorescence lifetime imaging (FLIM), and with advanced microscopy platforms using confocal, pulsed, or modulated illumination.

F&#246;rster resonance energy transfer (FRET) imaging is a powerful tool for real-time cell biology studies. Here a method for FRET imaging cells in physiologic three-dimensional (3D) hydrogel microenvironments using conventional epifluorescence microscopy is presented. An analysis for ratiometric FRET probes that yields linear ratios over the activation range is described.

Imaging of F&#246;rster resonance energy transfer (FRET)&#160;is a powerful tool for examining cell biology in real-time. Studies utilizing FRET commonly ...

Three-dimensional Reconstruction of the Vascular Architecture of the Passive CLARITY-cleared Mouse Ovary

The ovary is the main organ of the female reproductive system and is essential for the production of female gametes and for controlling the endocrine system, but the complex structural relationships and three-dimensional (3D) vasculature architectures of the ovary are not well described. In order to visualize the 3D connections and architecture of blood vessels in the intact ovary, the first important step is to make the ovary optically clear. In order to avoid tissue shrinkage, we used the hydrogel fixation-based passive CLARITY (Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging/ Immunostaining/In situ-hybridization-compatible Tissue Hydrogel) protocol method to clear an intact ovary. Immunostaining, advanced multiphoton confocal microscopy, and 3D image-reconstructions were then used for the visualization of ovarian vessels and follicular capillaries. Using this approach, we showed a significant positive correlation (P &#60;0.01) between the length of the follicular capillaries and volume of the follicular wall.

Here we present an adaptation of the passive CLARITY and 3D reconstruction method for visualization of the ovarian vasculature and follicular capillaries in intact mouse ovaries.

The ovary is the main organ of the female reproductive system and is essential for the production of female gametes and for controlling the endocrine ...

Single-Cell Resolution Three-Dimensional Imaging of Intact Organoids

Organoid technology, in vitro 3D culturing of miniature tissue, has opened a new experimental window for cellular processes that govern organ development and function as well as disease. Fluorescence microscopy has played a major role in characterizing their cellular composition in detail and demonstrating their similarity to the tissue they originate from. In this article, we present a comprehensive protocol for high-resolution 3D imaging of whole organoids upon immunofluorescent labeling. This method is widely applicable for imaging of organoids differing in origin, size and shape. Thus far we have applied the method to airway, colon, kidney, and liver organoids derived from healthy human tissue, as well as human breast tumor organoids and mouse mammary gland organoids. We use an optical clearing agent, FUnGI, which enables the acquisition of whole 3D organoids with the opportunity for single-cell quantification of markers. This three-day protocol from organoid harvesting to image analysis is optimized for 3D imaging using confocal microscopy.

The entire 3D structure and cellular content of organoids, as well as their phenotypic resemblance to the original tissue can be captured using the single-cell resolution 3D imaging protocol described here. This protocol can be applied to a wide range of organoids varying in origin, size and shape.

Organoid technology, in vitro 3D culturing of miniature tissue, has opened a new experimental window for cellular processes that govern organ development ...

Evaluating the Angiogenetic Properties of Ovarian Cancer Stem-Like Cells using the Three-Dimensional Co-Culture System, NICO-1

Cancer stem cells (CSCs) reside in a supportive niche, constituting a microenvironment comprised of adjacent stromal cells, vessels, and extracellular matrix. The ability of CSCs to participate in the development of endothelium constitutes an important characteristic that directly contributes to the general understanding of the mechanisms of tumorigenesis and tumor metastasis. The purpose of this work is to establish a reproducible methodology to investigate the tumor-initiation capability of ovarian cancer stem cells (OCSCs). Herein, we examined the neovascularization mechanism between endothelial cells and OCSCs along with the morphological changes of endothelial cells using the in vitro co-culture model NICO-1. This protocol allows visualization of the neovascularization step surrounding the OCSCs in a time course manner. The technique can provide insight regarding the angiogenetic properties of OCSCs in tumor metastasis.

Ovarian cancer stem cells (OCSC) are responsible for cancer initiation, recurrence, therapeutic resistance, and metastasis. The OCSC vascular niche is considered to promote self-renewal of OCSCs, leading to chemoresistance. This protocol provides the basis for establishing a reproducible OCSC vascular niche model in vitro.

Cancer stem cells (CSCs) reside in a supportive niche, constituting a microenvironment comprised of adjacent stromal cells, vessels, and extracellular ...

3D Culturing of Organoids from the Intestinal Villi Epithelium Undergoing Dedifferentiation

Clonogenicity of organoids from the intestinal epithelium is attributed to the presence of stem cells therein. The mouse small intestinal epithelium is compartmentalized into crypts and villi: the stem and proliferating cells are confined to the crypts, whereas the villi epithelium contains only differentiated cells. Hence, the normal intestinal crypts, but not the villi, can give rise to organoids in 3D cultures. The procedure described here is applicable only to villus epithelium undergoing dedifferentiation leading to stemness. The method described uses the Smad4-loss-of-function:&#946;-catenin gain-of-function (Smad4KO:&#946;-cateninGOF) conditional mutant mouse. The mutation causes the intestinal villi to dedifferentiate and generate stem cells in the villi. Intestinal villi undergoing dedifferentiation are scraped off the intestine using glass slides, placed in a 70 &#181;m strainer and washed several times to filter out any loose cells or crypts prior to plating in BME-R1 matrix to determine their organoid-forming potential. Two main criteria were used to ensure that the resulting organoids were developed from the dedifferentiating villus compartment and not from the crypts: 1) microscopically evaluating&#160;the isolated villi to ensure absence of any tethered crypts, both before and after plating in the 3D matrix, and 2) monitoring the time course of organoid development from the villi. Organoid initiation from the villi occurs only two to five days after plating and appears irregularly shaped, whereas the crypt-derived organoids from the same intestinal epithelium are apparent within sixteen hours of plating and appear spherical. The limitation of the method, however, is that the number of organoids formed, and the time required for organoid initiation from the villi vary depending on the degree of dedifferentiation. Hence, depending upon the specificity of the mutation or the insult causing the dedifferentiation, the optimal stage at which villi can be harvested to assay their organoid forming potential, must be determined empirically.

The procedure describes isolation of the villi from the mouse intestinal epithelium undergoing dedifferentiation to determine their organoid forming potential.

Clonogenicity of organoids from the intestinal epithelium is attributed to the presence of stem cells therein. The mouse small intestinal epithelium is ...

Construction of a Realistic, Whole-Body, Three-Dimensional Equine Skeletal Model using Computed Tomography Data

Therapies based upon whole-body biomechanical assessments are successful for injury prevention and rehabilitation in human athletes. Similar approaches have rarely been used to study equine athletic injury. Degenerative osteoarthritis caused by mechanical stress can originate from chronic postural dysfunction, which, because the primary dysfunction is often distant from the site of tissue injury, is best identified through modeling whole-body biomechanics. To characterize whole-body equine kinematics, a realistic skeletal model of a horse was created from equine computed tomography (CT) data that can be used for functional anatomical and biomechanical modeling. Equine CT data were reconstructed into individual three-dimensional (3D) data sets (i.e., bones) using 3D visualization software and assembled into a complete 3D skeletal model. The model was then rigged and animated using 3D animation and modeling software. The resulting 3D skeletal model can be used to characterize equine postures associated with degenerative tissue changes as well as to identify postures that reduce mechanical stress at the sites of tissue injury. In addition, when animated into 4D, the model can be used to demonstrate unhealthy and healthy skeletal movements and can be used to develop preventative and rehabilitative individualized therapies for horses with degenerative lamenesses. Although the model will soon be available for download, it is currently in a format that requires access to the 3D animation and modeling software, which has quite a learning curve for new users. This protocol will guide users in (1) developing such a model for any organism of interest and (2) using this specific equine model for their own research questions.

The purpose of this protocol is to describe the method of creation of a realistic, whole-body, skeletal model of a horse that can be used for functional anatomical and biomechanical modeling to characterize whole-body mechanics.

Therapies based upon whole-body biomechanical assessments are successful for injury prevention and rehabilitation in human athletes. Similar approaches ...