This work was funded by the KAMIN program of the Israeli Trade and Labor Ministry and by Qrons Inc., 777 Brickell Avenue Miami, FL 33131, US.

The use of animals in this protocol was approved by the National Animal Care and Use Committee.
NOTE: Calcium carbonated coral skeletons should be used in the crystalline form of aragonite. The coral types tested so far for neural cultures are Porites Lutea, Stylophora Pistillata, and Trachyphyllia Geoffroyi. The skeletons can be purchased whole or ground.
1. Cleaning the coral skeleton pieces
CAUTION: The following steps should be performed in a chemical hood at room temperature, because the solutions described below are hazardous and may cause burns and irritations.
<ol>
	<li>Use a hammer to break the coral skeleton and divide it into 0.5&#8211;2 cm fragments. In order to dissolve organic and non-organic residues, soak the coral skeleton fragments in 10% sodium hypochlorite solution for 10 min, then wash once with double distilled water (DDW).</li>
	<li>To remove the remaining organic residues, soak the fragments in a 1M NaOH solution for 5 min, then wash once with DDW. Continue with the removal of the organic deposits by soaking the fragments in 30% H2O2 solution for 10 min, then wash the fragments 3x with DDW.</li>
	<li>Remove as much excess DDW as possible and leave the coral fragments in the hood to dry (1&#8211;8 h).</li>
</ol>
2. Cleaning the glass coverslips
<ol>
	<li>Transfer the glass coverslips into a 100 mm glass Petri dish. Add 10 mL of 95% ethanol for 15 min.</li>
	<li>Remove the ethanol and wash with DDW 3x, waiting 10 min between each wash. Place the dish on an 80 &#176;C pre-warmed heating plate until the DDW evaporates. Gently stir the coverslips within the plate several times while drying to keep them from sticking to each other.</li>
	<li>Autoclave the coverslips.</li>
</ol>
3. Preparation of coral skeleton grains
<ol>
	<li>Grind the coral skeleton fragments using a mortar and pestle (manual grinding) until complete breakdown. The outcome is a mixture of grains with sizes ranging from 20 &#181;m&#8211;200 &#181;m.</li>
	<li>Alternatively, grind the coral skeleton fragments using an electrical grinding machine at a velocity of 1,000 rpm for 30 s (blade length = 6 cm; width = 0.5 cm&#8211;1.0 cm). The resulting grain size is similar to the size range produced by the manual grinding.</li>
</ol>
4. Purification of grains of a specific size range
NOTE: If control over the size of the grains in a matrix is desired, use the following filtration-based grain purification procedure.
<ol>
	<li>Transfer the grains onto a manual or electrical 40 &#181;m filter mesh strainer.</li>
	<li>Divide the grains into two specific ranges by sieving the grains through the strainer (if using an electrical strainer, the following conditions are recommended: shaking = 600 amplitudes/min, bouncing = 6/s). This procedure produces two groups of grain sizes, one &#60;40 &#181;m, and the second &#62;40 &#181;m. 
	NOTE: The two sizes can be determined by using strainers of varying meshes. If a more restricted range is desired, then re-sieve each group from step 4.2 through strainers with different meshes.</li>
	<li>Autoclave the grains.</li>
</ol>
5. Preparation of coral grain-coated dishes or coverslips
<ol>
	<li>For coating flasks, plates, or Petri dishes 
	NOTE: The following steps should be performed under sterile conditions.
	<ol>
		<li>Add the coral skeleton grains into a 20 &#181;g/mL poly-D-lysine (PDL) solution dissolved in Hanks&#39; solution. The concentration recommended is 5 mg/10 mg of grains per 1 mL PDL solution.</li>
		<li>Pour the solution into flasks and dishes (approximately 2 mL/25 cm2) and incubate overnight at 4 &#176;C. The grains sink and attach to the bottom.</li>
		<li>The next day, wash the flasks and dishes once with sterile DDW. Let the flasks and dishes dry in the hood. 
		NOTE: It is preferable to use freshly coated flasks and dishes. The coated flasks and dishes can be used up to a week after coating if preserved at 4 &#176;C. However, the effectiveness of the matrix may be compromised.</li>
	</ol>
	</li>
	<li>Coating glass coverslips (12 mm diameter, 0.17 mm thickness)
	<ol>
		<li>Add coral skeleton grains to DDW at any desired concentration. The common densities used for neural cells are 5 mg/mL and 10 mg/mL. Pour 40 &#181;L of the grain solution onto the center of the coverslip (Figure 4).</li>
		<li>Place the coverslips on a heating plate prewarmed to 80 &#176;C and wait for complete evaporation (usually 15 min). Under these conditions, the grains adhere to the coverslip. Autoclave the coated coverslips. Store in sterile conditions.</li>
		<li>A day before culture, place the coverslips on the lid of a sterile 24 well plate. Add 100 &#181;L of 20 &#181;g/mL PDL solution to each coverslip. Use the tip to ensure that the liquid covers the entire grain region and the rest of the coverslip surface.</li>
		<li>Cover the lid with the bottom of the plate. Wrap the sides of the plates with paraffin film. Incubate overnight at 4 &#176;C.</li>
		<li>The next day, wash once with DDW and dry in the hood. 
		NOTE: It is preferable to use freshly coated coverslips.</li>
	</ol>
	</li>
</ol>
6. Cultivation of hippocampal dissociated cells on coral skeleton grain-coated glass coverslips
NOTE: The method for hippocampal dissociated cell culture was modified from previously published procedures24,27. The preparation of the culture is described for four rat pups. The expected yield from each hippocampus is 1&#8211;1.5 x 106 cells.
<ol>
	<li>Setup
	<ol>
		<li>Prepare an empty 60 mm Petri dish. Pour 2 mL of minimal essential medium (MEM) into a 60 mm Petri dish and put on ice.</li>
		<li>Pour 2 mL of MEM into a 35 mm dish and put it in an incubator at 37 &#176;C, 5&#8211;10% CO2. Pour 2 mL of MEM into a 15 mL tube and keep it at 4 &#176;C.</li>
		<li>Pour 1 mL of First Day Medium into a 15 mL tube and put it in an incubator at 37 &#176;C, 5-10% CO2. 
		NOTE: The composition of the First Day Medium is described in the Table of Materials.</li>
		<li>Thaw 200 &#181;L of a 2.5% trypsin solution and incubate at 37 &#176;C.</li>
		<li>Prepare three glass Pasteur pipettes with 0.5 mm, 0.75 mm, and 1.0 mm diameter heads using a flame.</li>
		<li>Prepare adequate surgical tools: one big scissor, two small scissors, one big tweezer, four small tweezers, and one scalpel.</li>
		<li>Prepare a stereomicroscope in the hood.</li>
	</ol>
	</li>
	<li>Using a big scissor, sacrifice 0&#8211;3 day old Sprague Dawley rat pups by separating their heads from their bodies into an empty 60 mm Petri dish (step 6.1.1). Dispose of the bodies.</li>
	<li>Pick up the head by holding it from the mouth with a big tweezer. Cut the skin covering the skull with a small scissor.</li>
	<li>With a clean scissor, enter beneath the skull (on the side of the cerebellum) and cut the skull to the right and to the left to enable its removal. Using a small tweezer, peel the skull off from the brain.</li>
	<li>With a clean tweezer, separate the brain from the head and put it in a 60 mm Petri dish containing 2 mL of cold MEM (see step 6.1.2).</li>
	<li>Using two small tweezers, dissect the hippocampi under a stereomicroscope. Transfer the hippocampi into the prepared 35 mm dish (from step 6.1.2) containing MEM. 
	NOTE: Steps 6.3&#8211;6.6&#160;should be repeated for each pup.</li>
	<li>Cut the hippocampi to approximately 1 mm slices using the scalpel. Add 200 &#181;L of the trypsin solution (diluted to a final concentration of 0.25%). Mix gently and incubate at 37 &#176;C, 5&#8211;10% CO2 for 30 min.</li>
	<li>After incubation, add 2 mL of cold MEM (step 6.1.2) to the dish to deactivate the trypsin. Transfer the trypsinized tissue to a 15 mL tube containing 1 mL First Day Medium preheated to 37 &#176;C (step 6.1.3) using the glass Pasteur pipette with the largest diameter (step 6.1.5). 
	NOTE: The best medium to tissue (v:v) ratio for further processing of the tissue is 1 mL/8 hippocampi. Avoid transferring the medium with the tissue as much as possible.</li>
	<li>Triturate the tissue by passing it through the largest diameter glass pipette 10&#8211;15 times. Then, repeat this process using the glass pipette with the medium diameter. Continue triturating with the smallest diameter pipette. Avoid bubbling to reduce cell death.</li>
	<li>Let the remaining&#160;tissue pieces sink for 2&#8211;5 min, then transfer the supernatant into another 15 mL tube. Count the cells using a hemocytometer. 
	NOTE: The preferable cell density is 200,000&#8211;400,000 cells/mL. Use First Day Medium to dilute or centrifugation at 470 x g to concentrate the cells.</li>
	<li>Seed 100 &#181;L of cells on each glass coverslip. While seeding, make sure to cover the entire coverslip with cells. Incubate at 37 &#176;C, 5&#8211;10% CO2.</li>
	<li>The next day, add 500 &#181;L of Neuronal Growth Medium to the wells. 
	NOTE: The composition of the Neuronal Growth Medium is described in the Table of Materials.</li>
	<li>Gently transfer each coverslip to its appropriate well using a tweezer while removing the First Day Medium by tilting the coverslip.&#160;Suction the remaining media from the lid.</li>
	<li>Incubate at 37 &#176;C, 5&#8211;10% CO2. Avoid replacing the medium throughout incubation.&#160;Cultures can be maintained under these conditions up to 1 month. The major concern is humidity. Therefore, make sure to maintain the incubator maximally humidified.</li>
</ol>

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The authors declare that they have no competing financial interests.

The technique presented here describes a way to improve the maintenance and functionality of neural cells in culture. This is achieved by adhering the cells to a matrix made of coral skeleton grains that nurtures the cells and promotes their growth and activity. Using this technique increases the capacity of the neural culture model to mimic the cells&#39; environment in the brain.
The introduction of the matrix as a culture substrate has several advantages over other substrates used in classi...

Cultures of dissociated neural cells, in this case hippocampal cells, are a valuable experimental model for studying neural growth and function by providing high cell isolation and accessibility1,2,3. This type of culture is frequently used in neuroscience, drug development, and tissue engineering due to the large amount of information that can be collected, such as rates of growth and viability, neurotoxicity, neurite outgrowth and networking, synaptic connectivity and plasticity, morphological modifications, neurites organization and wiring, etc.1,4,5,6,7.
Despite the significance of the cultures, the cultivated cells are usually forced to grow on glass coverslips in a two-dimensional monolayer. These strict environmental modifications significantly decrease the ability of neural cells to survive over time, because glass coverslips are non-nurturing substrates with a low adhesion strength, exhibiting a lower capacity to support cell growth8,9,10,11.
Because cultivated neural cells are forced to grow in challenging conditions, an essential approach to enhance their survival would be to imitate their natural environment as much as possible12,13. This could be achieved by using biomaterials that will act as matrices and mimic the extracellular matrix of the cells, enabling them to form a tissue-like structure and assist in their nourishment14.
The use of biomaterials is a promising approach in improving cell cultures, because they act as biocompatible scaffolds, providing mechanical stability and enhancing a variety of cell properties, including adhesion, survival, proliferation, migration, morphogenesis, and differentiation15,16,17. Several types of biomaterials are used to improve the conditions of the cells in vitro. Among them are biopolymers, or biological components that are usually part of the extracellular matrix of the cells. These biomaterials are mostly used as a form of polymerized coating agents or hydrogels18,19,20. On the one hand, the matrices mentioned above give the cells a familiar 3D environment to grow in, encourage their adhesion to the dish, and give them mechanical support21,22. On the other hand, their polymerized form and the confinement of the cells within hydrogels disturbs the access of the cells to nurturing components present in the growth media and also makes the follow up of the cells by microscopic methods more difficult23.
Coral exoskeletons are biological marine-originated matrices. They are made of calcium carbonate, have mechanical stability, and are biodegradable. Previous studies using the coral skeleton as a matrix for growing neural cells in culture have shown much greater adhesion, compared to glass coverslips24,25. In addition, neural cells grown on coral skeleton demonstrated their capability to intake the calcium the skeleton is composed of, which protects the neural cells in conditions of nutrient deprivation26. Moreover, the coral skeleton is a supportive and nurturing matrix that increases the survival of neural cells, encourages the formation of neural networks, elevates the rate of synaptic connectivity, and enables the formation of tissue-like structures27,28. Recent studies have also shown that the surface topography of the coral skeleton matrix plays a crucial role in the distribution and activation of glial cells8,29. Also, coral skeleton is effective as a matrix for cultivation of other cell types, such as osteocytes30,31, hepatocytes, and cardiomyocytes in culture (unpublished data).
Hence, coral skeleton is a promising matrix for cultivation of cells in vitro. Thus, the protocol detailed below describes the technique of cultivating neural cells on coral skeleton for producing more stable and prosperous neural cultures than those achieved by existing methods. This protocol may also be useful for cultivation of cardiomyocytes, hepatocytes, and other cell types.

<table>
	<tbody>
		<tr>
			<td>24-well plates</td>
			<td>Greiner</td>
			<td>#60-662160</td>
			<td></td>
		</tr>
		<tr>
			<td>B-27</td>
			<td>Gibco</td>
			<td>#17504-044</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA)</td>
			<td>Sigma</td>
			<td>#A4503</td>
			<td></td>
		</tr>
		<tr>
			<td>D &#8211; glucose</td>
			<td>Sigma</td>
			<td>#G8769</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Minimal Essential Eagle (DMEM)</td>
			<td>Sigma</td>
			<td>#D5796</td>
			<td></td>
		</tr>
		<tr>
			<td>Electrical sieve</td>
			<td>Ari Levy</td>
			<td>#3700</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serun (FBS)</td>
			<td>Biological Industries</td>
			<td>#04-007-1A</td>
			<td></td>
		</tr>
		<tr>
			<td>First Day Medium</td>
			<td></td>
			<td></td>
			<td>85.1% Minimum Essential Eagle&#8217;s medium (MEM), 11.5% heat-inactivated fetal bovine serum, 1.2% L-Glutamine and 2.2% D-Glucose.</td>
		</tr>
		<tr>
			<td>Flasks</td>
			<td>Greiner</td>
			<td>#60-690160</td>
			<td>25cm^2, Tissue culture treated</td>
		</tr>
		<tr>
			<td>Fluoro-deoxy-uridine</td>
			<td>Sigma</td>
			<td>#F0503</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Coverslips</td>
			<td>Menzel-Glaser</td>
			<td>#BNCB00120RA1</td>
			<td></td>
		</tr>
		<tr>
			<td>H2O2</td>
			<td>Romical</td>
			<td>#007130-72-19</td>
			<td>Hazardous</td>
		</tr>
		<tr>
			<td>Ham&#39;s F-12 Nutrient Mixture</td>
			<td>Sigma</td>
			<td>#N4888</td>
			<td></td>
		</tr>
		<tr>
			<td>HANK&#39;S solution</td>
			<td>Sigma</td>
			<td>#H6648</td>
			<td></td>
		</tr>
		<tr>
			<td>Kynurenic acid</td>
			<td>Sigma</td>
			<td>#K3375</td>
			<td></td>
		</tr>
		<tr>
			<td>L - glutamine</td>
			<td>Sigma</td>
			<td>#G7513</td>
			<td></td>
		</tr>
		<tr>
			<td>Manual strainer (40&#181;m)</td>
			<td>VWR</td>
			<td>#10199-654</td>
			<td></td>
		</tr>
		<tr>
			<td>Minimun Essential Eagle (MEM)</td>
			<td>Sigma</td>
			<td>#M2279</td>
			<td></td>
		</tr>
		<tr>
			<td>Mortar and pestle</td>
			<td>De-Groot</td>
			<td>4-P090</td>
			<td></td>
		</tr>
		<tr>
			<td>NaClO (Sodium Hypochlorite)</td>
			<td>Sigma</td>
			<td>#425044</td>
			<td>Hazardous</td>
		</tr>
		<tr>
			<td>NaOH</td>
			<td>Sigma</td>
			<td>#S8045</td>
			<td>Hazardous</td>
		</tr>
		<tr>
			<td>Neuronal Growth Medium</td>
			<td></td>
			<td></td>
			<td>45% MEM, 40% Dulbecco&#39;s modified eagle&#39;s medium (DMEM), 10% Nutrient mixture F-12 Ham, 0.25% (w/v) bovine serum albumin (BSA), 0.75% D-glucose, 0.25% L-Glutamine, 0.5% B-27 supplement, 0.1% kynurenic acid, 0.01% of 70 % uridine and 30% fluoro-deoxy-uridine.</td>
		</tr>
		<tr>
			<td>Petri dish</td>
			<td>Greiner</td>
			<td>#60-628160, #60-627160</td>
			<td>60mm, 35mm, respectively.</td>
		</tr>
		<tr>
			<td>Poly D &#8211; Lysine</td>
			<td>Sigma</td>
			<td>#P7280</td>
			<td></td>
		</tr>
		<tr>
			<td>Smart Dentin Grinder</td>
			<td>KometaBio</td>
			<td>#GR101</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin</td>
			<td>Gibco</td>
			<td>#15-090-046</td>
			<td></td>
		</tr>
		<tr>
			<td>Uridine</td>
			<td>Sigma</td>
			<td>#U3750</td>
			<td></td>
		</tr>
	</tbody>
</table>

increasing durability of dissociated neural cell cultures using biologically active coralline matrix

Cultures of dissociated hippocampal neuronal and glial cells are a valuable experimental model for studying neural growth and function by providing high cell isolation and a controlled environment. However, the survival of hippocampal cells in vitro is compromised: most cells die during the first week of culture. It is therefore of great importance to identify ways to increase the durability of neural cells in culture.
Calcium carbonate in the form of crystalline aragonite derived from the skeleton of corals can be used as a superior, active matrix for neural cultures. By nurturing, protecting, and activating glial cells, the coral skeleton enhances the survival and growth of these cells in vitro better than other matrices.
This protocol describes a method for cultivating hippocampal cells on a coralline matrix. This matrix is generated by attaching grains of coral skeletons to culture dishes, flasks, and glass coverslips. The grains assist in improving the environment of the cells by introducing them to a fine three-dimensional (3D) environment to grow on and to form tissue-like structures. The 3D environment introduced by the coral skeleton can be optimized for the cells by grinding, which enables control over the size and density of the grains (i.e., the matrix roughness), a property that has been found to influence glial cells activity. Moreover, the use of grains makes the observation and analysis of the cultures easier, especially when using light microscopy. Hence, the protocol includes procedures for generation and optimization of the coralline matrix as a tool to improve the maintenance and functionality of neural cells in vitro.

In order to prepare the coral skeleton matrix, the entire coral skeleton (Figure 1A) was broken into 0.5&#8211;2 cm fragments using a hammer (Figure 1B) and thoroughly cleaned from organic residues through three steps (step 1 in the protocol) using 10% hypochlorite solution, 1M NaOH solution, and 30% H2O2 solution (Figure 1C). Coral fragments were well-cleaned when the skeleton color changed from brown (<strong...

Watch this Scientific Journal Video about Increasing Durability of Dissociated Neural Cell Cultures Using Biologically Active Coralline Matrix at JoVE.com

Increasing Durability of Dissociated Neural Cell Cultures Using Biologically Active Coralline Matrix

Dissociated hippocampal cell culture is a pivotal experimental tool in neuroscience. Neural cell survival and function in culture is enhanced when coralline skeletons are used as matrices, due to their neuroprotective and neuromodulative roles. Hence, neural cells grown on coralline matrix show higher durability, and thereby are more adequate for culturing.

Cultures of dissociated hippocampal neuronal and glial cells are a valuable experimental model for studying neural growth and function by providing high ...

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Bioengineering

1.2K Views.  Ariel University. Dissociated hippocampal cell culture is a pivotal experimental tool in neuroscience. Neural cell survival and function in culture is enhanced when coralline skeletons are used as matrices, due to their neuroprotective and neuromodulative roles. Hence, neural cells grown on coralline matrix show higher durability, and thereby are more adequate for culturing.

Video: Increasing Durability of Dissociated Neural Cell Cultures Using Biologically Active Coralline Matrix

Fabrication of Biologically Derived Injectable Materials for Myocardial Tissue Engineering

This protocol provides methods for the preparation of an injectable extracellular matrix (ECM) gel for myocardial tissue engineering applications. Briefly, decellularized tissue is lyophilized, milled, enzymatically digested, and then brought to physiological pH. The lyophilization removes all water content from the tissue, resulting in dry ECM that can be ground into a fine powder with a small mill. After milling, the ECM powder is digested with pepsin to form an injectable matrix. After adjustment to pH 7.4, the liquid matrix material can be injected into the myocardium. Results of previous characterization assays have shown that matrix gels produced from decellularized pericardial and myocardial tissue retain native ECM components, including diverse proteins, peptides and glycosaminoglycans. Given the use of this material for tissue engineering, in vivo characterization is especially useful; here, a method for performing an intramural injection into the left ventricular (LV) free wall is presented as a means of analyzing the host response to the matrix gel in a small animal model. Access to the chest cavity is gained through the diaphragm and the injection is made slightly above the apex in the LV free wall. The biologically derived scaffold can be visualized by biotin-labeling before injection and then staining tissue sections with a horse radish peroxidase-conjugated neutravidin and visualizing via diaminobenzidine (DAB) staining. Analysis of the injection region can also be done with histological and immunohistochemical staining. In this way, the previously examined pericardial and myocardial matrix gels were shown to form fibrous, porous networks and promote vessel formation within the injection region.

Methods for preparing an injectable matrix gel from decellularized tissue and injecting it into rat myocardium in vivo are described.

This protocol provides methods for the preparation of an injectable extracellular matrix (ECM) gel for myocardial tissue engineering applications.  ...

Shape Memory Polymers for Active Cell Culture

Shape memory polymers (SMPs) are a class of "smart" materials that have the ability to change from a fixed, temporary shape to a pre-determined permanent shape upon the application of a stimulus such as heat1-5. In a typical shape memory cycle, the SMP is first deformed at an elevated temperature that is higher than its transition temperature, Ttrans [either the melting temperature (Tm) or the glass transition temperature (Tg)]. The deformation is elastic in nature and mainly leads to a reduction in conformational entropy of the constituent network chains (following the rubber elasticity theory). The deformed SMP is then cooled to a temperature below its Ttrans while maintaining the external strain or stress constant. During cooling, the material transitions to a more rigid state (semi-crystalline or glassy), which kinetically traps or "freezes" the material in this low-entropy state leading to macroscopic shape fixing. Shape recovery is triggered by continuously heating the material through Ttrans under a stress-free (unconstrained) condition. By allowing the network chains (with regained mobility) to relax to their thermodynamically favored, maximal-entropy state, the material changes from the temporary shape to the permanent shape.
Cells are capable of surveying the mechanical properties of their surrounding environment6. The mechanisms through which mechanical interactions between cells and their physical environment control cell behavior are areas of active research. Substrates of defined topography have emerged as powerful tools in the investigation of these mechanisms. Mesoscale, microscale, and nanoscale patterns of substrate topography have been shown to direct cell alignment, cell adhesion, and cell traction forces7-14. These findings have underscored the potential for substrate topography to control and assay the mechanical interactions between cells and their physical environment during cell culture, but the substrates used to date have generally been passive and could not be programmed to change significantly during culture. This physical stasis has limited the potential of topographic substrates to control cells in culture.
Here, active cell culture (ACC) SMP substrates are introduced that employ surface shape memory to provide programmed control of substrate topography and deformation. These substrates demonstrate the ability to transition from a temporary grooved topography to a second, nearly flat memorized topography. This change in topography can be used to control cell behavior under standard cell culture conditions.

A method for developing cell culture substrates with the ability to change topography during culture is described. The method makes use of smart materials known as shape memory polymers that have the ability to memorize a permanent shape. This concept is adaptable to a wide range of materials and applications.

Shape memory polymers (SMPs) are a class of "smart" materials that have the ability to change from a fixed, temporary shape to a pre-determined permanent ...

Increasing cDNA Yields from Single-cell Quantities of mRNA in Standard Laboratory Reverse Transcriptase Reactions using Acoustic Microstreaming

Correlating gene expression with cell behavior is ideally done at the single-cell level. However, this is not easily achieved because the small amount of labile mRNA present in a single cell (1-5% of 1-50pg total RNA, or 0.01-2.5pg mRNA, per cell 1) mostly degrades before it can be reverse transcribed into a stable cDNA copy. For example, using standard laboratory reagents and hardware, only a small number of genes can be qualitatively assessed per cell 2. One way to increase the efficiency of standard laboratory reverse transcriptase (RT) reactions (i.e. standard reagents in microliter volumes) comprising single-cell amounts of mRNA would be to more rapidly mix the reagents so the mRNA can be converted to cDNA before it degrades. However this is not trivial because at microliter scales liquid flow is laminar, i.e. currently available methods of mixing (i.e. shaking, vortexing and trituration) fail to produce sufficient chaotic motion to effectively mix reagents. To solve this problem, micro-scale mixing techniques have to be used 3,4. A number of microfluidic-based mixing technologies have been developed which successfully increase RT reaction yields 5-8. However, microfluidics technologies require specialized hardware that is relatively expensive and not yet widely available. A cheaper, more convenient solution is desirable. The main objective of this study is to demonstrate how application of a novel "micromixing" technique to standard laboratory RT reactions comprising single-cell quantities of mRNA significantly increases their cDNA yields. We find cDNA yields increase by approximately 10-100-fold, which enables: (1) greater numbers of genes to be analyzed per cell; (2) more quantitative analysis of gene expression; and (3) better detection of low-abundance genes in single cells. The micromixing is based on acoustic microstreaming 9-12, a phenomenon where sound waves propagating around a small obstacle create a mean flow near the obstacle. We have developed an acoustic microstreaming-based device ("micromixer") with a key simplification; acoustic microstreaming can be achieved at audio frequencies by ensuring the system has a liquid-air interface with a small radius of curvature 13. The meniscus of a microliter volume of solution in a tube provides an appropriately small radius of curvature. The use of audio frequencies means that the hardware can be inexpensive and versatile 13, and nucleic acids and other biochemical reagents are not damaged like they can be with standard laboratory sonicators.

We describe a novel method for increasing cDNA yield from single-cell quantities of mRNA in otherwise standard laboratory reverse transcription reactions. The novelty resides in the use of a micromixer, which utilizes the phenomenon of acoustic microstreaming, to mix fluids at microliter scales more effectively than shaking, vortexing or trituration.

Correlating gene expression with cell behavior is ideally done at the single-cell level.  However, this is not easily achieved because the small amount of ...

Non-contact, Label-free Monitoring of Cells and Extracellular Matrix using Raman Spectroscopy

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, currently available routine methods require processing steps and alter sample integrity. Raman spectroscopy is a fast method that enables the measurement of biological samples without the need for further processing steps. This laser-based technology detects the inelastic scattering of monochromatic light.6 As every chemical vibration is assigned to a specific Raman band (wavenumber in cm-1), each biological sample features a typical spectral pattern due to their inherent biochemical composition.7-9 Within Raman spectra, the peak intensities correlate with the amount of the present molecular bonds.1 Similarities and differences of the spectral data sets can be detected by employing a multivariate analysis (e.g. principal component analysis (PCA)).10
Here, we perform Raman spectroscopy of living cells and native tissues. Cells are either seeded on glass bottom dishes or kept in suspension under normal cell culture conditions (37 &deg;C, 5% CO2) before measurement. Native tissues are dissected and stored in phosphate buffered saline (PBS) at 4 &deg;C prior measurements. Depending on our experimental set up, we then either focused on the cell nucleus or extracellular matrix (ECM) proteins such as elastin and collagen. For all studies, a minimum of 30 cells or 30 random points of interest within the ECM are measured. Data processing steps included background subtraction and normalization.

Raman spectroscopy is a suitable technique for the non-contact, label-free analysis of living cells, tissue-engineered constructs and native tissues. Source-specific spectral fingerprints can be generated and analyzed using multivariate analysis.

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, ...

Shock Wave Application to Cell Cultures

Shock waves nowadays are well known for their regenerative effects. Basic research findings showed that shock waves do cause a biological stimulus to target cells or tissue without any subsequent damage. Therefore, in vitro experiments are of increasing interest. Various methods of applying shock waves onto cell cultures have been described. In general, all existing models focus on how to best apply shock waves onto cells.
However, this question remains: What happens to the waves after passing the cell culture? The difference of the acoustic impedance of the cell culture medium and the ambient air is that high, that more than 99% of shock waves get reflected! We therefore developed a model that mainly consists of a Plexiglas built container that allows the waves to propagate in water after passing the cell culture. This avoids cavitation effects as well as reflection of the waves that would otherwise disturb upcoming ones. With this model we are able to mimic in vivo conditions and thereby gain more and more knowledge about how the physical stimulus of shock waves gets translated into a biological cell signal (&#8220;mechanotransduction&#34;).

Shock waves nowadays are well known for their regenerative effects. Therefore in vitro experiments are of increasing interest. We therefore developed a model for in vitro shock wave trials (IVSWT) that enables us to mimic in vivo conditions thereby avoiding distracting physical effects.

Shock waves nowadays are well known for their regenerative effects. Basic research findings showed that shock waves do cause a biological stimulus to ...

Analysis of Cell Migration within a Three-dimensional Collagen Matrix

The ability to migrate is a hallmark of various cell types and plays a crucial role in several physiological processes, including&#160;embryonic development, wound healing, and immune responses. However, cell migration is also a key mechanism in cancer enabling these cancer cells to detach from the primary tumor to start metastatic spreading. Within the past years various cell migration assays have been developed to analyze the migratory behavior of different cell types. Because the locomotory behavior of cells markedly differs between a two-dimensional (2D) and three-dimensional (3D) environment it can be assumed that the analysis of the migration of cells that are embedded within a 3D environment would yield in more significant cell migration data. The advantage of the described 3D collagen matrix migration assay is that cells are embedded within a physiological 3D network of collagen fibers representing the major component of the extracellular matrix. Due to time-lapse video microscopy real cell migration is measured allowing the determination of several migration parameters as well as their alterations in response to pro-migratory factors or inhibitors. Various cell types could be analyzed using this technique, including lymphocytes/leukocytes, stem cells, and tumor cells. Likewise, also cell clusters or spheroids could be embedded within the collagen matrix concomitant with analysis of the emigration of single cells from the cell cluster/ spheroid into the collagen lattice. We conclude that the 3D collagen matrix migration assay is a versatile method to analyze the migration of cells within a physiological-like 3D environment.

Cell migration is a biological phenomenon that is involved in a plethora of physiological, such as wound healing and immune responses, and pathophysiological processes, like cancer. The 3D-collagen matrix migration assay is a versatile tool to analyze the migratory properties of different cell types within in a 3D physiological-like environment.

The ability to migrate is a hallmark of various cell types and plays a crucial role in several physiological processes, including&#160;embryonic ...

Improved 3D Hydrogel Cultures of Primary Glial Cells for In Vitro Modelling of Neuroinflammation

In the central nervous system, numerous acute injuries and neurodegenerative disorders, as well as implanted devices or biomaterials engineered to enhance function result in the same outcome: excess inflammation leads to gliosis, cytotoxicity, and/or formation of a glial scar that collectively exacerbate injury or prevent healthy recovery. With the intent of creating a system to model glial scar formation and study inflammatory processes, we have generated a 3D cell scaffold capable of housing primary cultured glial cells: microglia that regulate the foreign body response and initiate the inflammatory event, astrocytes that respond to form a fibrous scar, and oligodendrocytes that are typically vulnerable to inflammatory injury. The present work provides a detailed step-by-step method for the fabrication, culture, and microscopic characterization of a hyaluronic acid-based 3D hydrogel scaffold with encapsulated rat brain-derived glial cells. Further, protocols for characterization of cell encapsulation and the hydrogel scaffold by confocal immunofluorescence and scanning electron microscopy are demonstrated, as well as the capacity to modify the scaffold with bioactive substrates, with incorporation of a commercial basal lamina mixture to improved cell integration.

Improved 3D Hydrogel Cultures of Primary Glial Cells for In Vitro Modelling of Neuroinflammation

Herein, we present a protocol for the 3D culture of rat brain-derived glia cells, including astrocytes, microglia, and oligodendrocytes. We demonstrate primary cell culture, methacrylated hyaluronic acid (HAMA) hydrogel synthesis, HAMAphoto-polymerization and cell encapsulation, and sample processing for confocal and scanning electron microscopic imaging.

In the central nervous system, numerous acute injuries and neurodegenerative disorders, as well as implanted devices or biomaterials engineered to enhance ...

A Novel Tenorrhaphy Suture Technique with Tissue Engineered Collagen Graft to Repair Large Tendon Defects

Surgical management of large tendon defects with tendon grafts is challenging, as there are a finite number of sites where donors can be readily identified and used. Currently, this gap is filled with tendon auto-, allo-, xeno-, or artificial grafts, but clinical methods to secure them are not necessarily translatable to animals because of the scale. In order to evaluate new biomaterials or study a tendon graft made up of collagen type 1, we have developed a modified suture technique to help maintain the engineered tendon in alignment with the tendon ends. Mechanical properties of these grafts are inferior to the native tendon. To incorporate engineered tendon into clinically relevant models of loaded repair, a strategy was adopted to offload the tissue engineered tendon graft and allow for the maturation and integration of the engineered tendon in vivo until a mechanically sound neo-tendon was formed. We describe this technique using incorporation of the collagen type 1 tissue engineered tendon construct.

In this paper, we present an in vitro and in situ protocol to repair a tendon gap of up to 1.5 cm by filling it with engineered collagen graft. This was performed by developing a modified suture technique to take the mechanical load until the graft matures into the host tissue.

Surgical management of large tendon defects with tendon grafts is challenging, as there are a finite number of sites where donors can be readily ...

Fabricating a Kidney Cortex Extracellular Matrix-Derived Hydrogel

Extracellular matrix (ECM) provides important biophysical and biochemical cues to maintain tissue homeostasis. Current synthetic hydrogels offer robust mechanical support for in vitro cell culture but lack the necessary protein and ligand composition to elicit physiological behavior from cells. This manuscript describes a fabrication method for a kidney cortex ECM-derived hydrogel with proper mechanical robustness and supportive biochemical composition. The hydrogel is fabricated by mechanically homogenizing and solubilizing decellularized human kidney cortex ECM. The matrix preserves native kidney cortex ECM protein ratios while also enabling gelation to physiological mechanical stiffnesses. The hydrogel serves as a substrate upon which kidney cortex-derived cells can be maintained under physiological conditions. Furthermore, the hydrogel composition can be manipulated to model a diseased environment which enables the future study of kidney diseases.

Here we present a protocol to fabricate a kidney cortex extracellular matrix-derived hydrogel to retain the native kidney extracellular matrix (ECM) structural and biochemical composition. The fabrication process and its applications are described. Finally, a perspective on using this hydrogel to support kidney-specific cellular and tissue regeneration and bioengineering is discussed.

Extracellular matrix (ECM) provides important biophysical and biochemical cues to maintain tissue homeostasis. Current synthetic hydrogels offer robust ...

Production of Extracellular Matrix Fibers via Sacrificial Hollow Fiber Membrane Cell Culture

Engineered scaffolds derived from extracellular matrix (ECM) have driven significant interest in medicine for their potential in expediting wound closure and healing. Extraction of extracellular matrix from fibrogenic cell cultures in vitro has potential for generation of ECM from human- and potentially patient-specific cell lines, minimizing the presence of xenogeneic epitopes which has hindered the clinical success of some existing ECM products. A significant challenge in in vitro production of ECM suitable for implantation is that ECM production by cell culture is typically of relatively low yield. In this work, protocols are described for the production of ECM by cells cultured within sacrificial hollow fiber membrane scaffolds. Hollow fiber membranes are cultured with fibroblast cell lines in a conventional cell medium and dissolved after cell culture to yield continuous threads of ECM. The resulting ECM fibers produced by this method can be decellularized and lyophilized, rendering it suitable for storage and implantation.

The goal of this protocol is production of whole extracellular matrix fibers targeted for wound repair which are suitable for preclinical evaluation as part of a regenerative scaffold implant. These fibers are produced by the culture of fibroblasts on hollow fiber membranes and extracted by dissolution of the membranes.

Engineered scaffolds derived from extracellular matrix (ECM) have driven significant interest in medicine for their potential in expediting wound closure ...