The authors would like to thank Baden-W&#252;rttemberg Stiftung for their financial support in the "Bioinspired Material Synthesis" framework (BioMatS-14).

1. Hydrogel Preparation
<ol>
	<li>Mix 200 mg of BSA with 1 mL of deionized H2O to create 20% (w/v) BSA stock (stock solution A).</li>
	<li>Mix 165 &#181;L of THPC solution (134 mg/mL) with 4.835 mL of deionized water to create THPC stock solution (stock solution B).</li>
	<li>Weigh 1 mg of KCSSGKSRGDS (1,111.1 g/mol) peptide (or an equivalent cell-adhesive peptide) and dilute it in 100 &#181;L of sterile H2O to obtain a 10 mg/mL solution (stock solution C). 
	NOTE: This step is optional and only needs to be included if the hydrogel is meant for cell culture application.</li>
	<li>Remove the bottom of a 96-well plate and replace it with removable plastic wrap. 
	NOTE: For the plates used here, the bottom can easily be removed by applying pressure to the bottom of each well; the thin plastic bottom will simply fall out.</li>
	<li>Mix 100 &#181;L of BSA stock solution (A) and 100 &#181;L of THPC stock solution (B) (optional: 4 &#181;L of stock solution C for functionalized, cell-adhesive hydrogels) in the 96-well plate to obtain 200 &#181;L of hydrogel. Mix the components by pipetting up and down at least 5 times to guarantee a uniform hydrogel after polymerization.</li>
	<li>Place the 96-well plate at room temperature (RT) for about 10 min until all hydrogels are properly polymerized.</li>
	<li>Carefully and slowly remove the plastic wrap from the bottom of the plate.</li>
	<li>Press the hydrogels out of the 96-well plate using a small stamp and transfer them to 1.5 mL tubes with sterile PBS, pH 7.4. 
	NOTE: The hydrogels have a cylindrical shape, with a diameter of approximately 4 mm and a height of 8 mm.</li>
	<li>Store the hydrogels in phosphate-buffered saline (PBS) at 4 &#176;C for up to several months.</li>
</ol>
2. Freeze-drying the Hydrogels
<ol>
	<li>Fill 1.5 mL tubes with 500 &#181;L of sterile deionized H2O and remove the cap. Transfer the hydrogels to the 1.5 mL reaction tubes using a spatula.</li>
	<li>Wrap the 1.5 mL tubes tightly with paraffin film-at least three layers for each vial. Use a needle to pierce small holes in the film to enable gas release from the tube.</li>
	<li>Proceed to one of the following steps:
	<ol>
		<li>Transfer the vial to liquid nitrogen solution for 5 min to guarantee that the water and hydrogel completely freeze. Immediately after removal from the liquid nitrogen, transfer the vials to the freeze dryer to prevent the material from thawing. 
		NOTE: This procedure results in pores about 10-15 &#181;m in radius.</li>
		<li>Keep the vials at -20 &#176;C overnight to slowly freeze the hydrogels. Immediately after removal from -20 &#176;C, transfer the vials to the freeze dryer to prevent the material from thawing. 
		NOTE: This procedure results in pores about 50-60 &#181;m in radius.</li>
	</ol>
	</li>
	<li>After 24 h (and the complete evaporation of the water in and around the hydrogel), thaw the material by removing it from the freeze dryer. 
	NOTE: The pore sizes can be analyzed with confocal laser scanning microscopy (step 5, &#34;Hydrogel Visualization&#34;).</li>
</ol>
3. Particle Leaching
<ol>
	<li>Prepare the hydrogel as described in steps 1.1-1.5.</li>
	<li>Directly after mixing the components, add NaCl until saturation occurs (36 mg/mL). Add salt until a salt crystal can be seen in the solution as a white precipitate.</li>
	<li>Transfer the 96-well plate to a shaker and shake until polymerization takes place (about 10 min). Remove the hydrogels from the plate, as described in steps 1.7-1.8.</li>
	<li>Incubate the hydrogels for at least 24 h at RT in sterile water on a shaker (20 rpm) to elute all salt from the hydrogel template. Store the hydrogels at 4 &#176;C in PBS for up to several months.</li>
</ol>
4. Channel Formation
<ol>
	<li>Prepare hydrogels as described in step 1. Remove a hydrogel from solution using a pincer, remove the excess water with highly absorbent paper, and place it on top of a block of dry ice.</li>
	<li>Freeze the hydrogel for 30 s and carefully remove it from the block. Do not damage the hydrogel; carefully use a spatula to scrape it off the block. Transfer the hydrogel to a 1.5 mL tube and dry it overnight at 37 &#176;C.</li>
</ol>
5. Hydrogel Visualization
<ol>
	<li>Prepare a rhodamine B stock solution by diluting 1 mg of rhodamine B in 10 mL of PBS. Prepare a serial dilution by diluting the rhodamine stock solution in PBS until a concentration of 0.001 mg/mL is reached (dilution factor: 100)</li>
	<li>Remove the hydrogel from the storage solution (e.g., from step 2.4.) and transfer it to a 1.5 mL tube with 1 mL of 0.01 mg/mL rhodamine B solution. Stain the hydrogel overnight in rhodamine B solution at RT.</li>
	<li>The next day, transfer the hydrogel that is to be visualized to 10 mL of PBS (pH 7.4) and wash for at least 3 h.</li>
	<li>Transfer the hydrogel onto a &#181;-slide 8 well and cover it with PBS.</li>
	<li>Cut small slices out of the hydrogel (about 0.5 mm in height) using a blade.</li>
	<li>Using a confocal laser scanning microscope, visualize the hydrogel at a wavelength of 514 nm (objective: EC Plan-Neofluar 40x/1.30 Oil DIC M27, plane scan mode: 514 nm, and zoom: 1X).</li>
</ol>
6. Cell Culture Feasibility
<ol>
	<li>Sterile-filter all stock solutions (A, B, and C) with a 0.45 &#181;m filter.</li>
	<li>Prepare the hydrogel as described in steps 1.1-1.4, including the cell-adhesive peptide prior to hydrogel polymerization.</li>
	<li>After mixing the components, immediately pipette the mixture into a &#181;-slide 8 well until the bottom is completely covered. 
	NOTE: This step must be performed quickly and directly after mixing the components, as polymerization takes place within minutes in the slide.</li>
	<li>Culture adhesion cells and passage to obtain a single-cell suspension using standard cell culture techniques. Transfer the cells into pre-warmed, sterile DMEM cell culture medium supplemented with fetal bovine serum (FBS, 10% (w/v)), penicillin-streptomycin (1% (w/v)), and nonessential amino acid solution (MEM, 1% (w/v)).</li>
	<li>Count the cells with a Neubauer counting chamber and carefully pipette 200 &#181;L of the desired number of cells (2 x 105 cells/cm2) onto the hydrogel surface.</li>
	<li>Cover the &#181;-slide 8 well with the lid and transfer it to an incubator (37 &#176;C, 5% CO2). Incubate for at least 4 h at 37 &#176;C.</li>
	<li>After at least 4 h of cellular attachment, wash the cells twice with 200 &#181;L of sterile cell culture PBS.</li>
	<li>Fix the cells with 200 &#181;L of 3.7% (v/v) formaldehyde for 10 min at RT and wash twice with PBS (~200 &#181;L). 
	NOTE: Wear appropriate personal protective equipment when handling formaldehyde.</li>
	<li>Permeabilize the cells with 200 &#181;L of 0.1 % Triton X for 5 min. Wash twice with PBS.</li>
	<li>Stain the cells with phalloidin-rhodamine by mixing 5 &#181;L of methanolic stock with 195 &#181;L of PBS and adding it to the cells at RT. Stain the cells for 20 min in the dark. Wash twice with PBS.</li>
	<li>Investigate the cell adhesion properties using a confocal microscope at 514 nm (objective: EC Plan-Neofluar 40x/1.30 Oil DIC M27, plane scan mode: 514 nm, and zoom: 1X). Analyze the images using appropriate software (see the Table of Materials).</li>
</ol>
7. Hydrogel Properties
<ol>
	<li>Swelling ratio.
	<ol>
		<li>Completely dry the hydrogels at 37 &#176;C for at least one day.</li>
		<li>Weigh each hydrogel and note the exact weight.</li>
		<li>Fill a 2 mL reaction tube with 1.5 mL of PBS.</li>
		<li>Transfer the hydrogel into this 2 mL reaction tube and completely immerse it in PBS.</li>
		<li>Leave the hydrogel for at least two days in PBS at RT to reach equilibrium with the PBS.</li>
		<li>After three days, remove the hydrogel from the solution and dry it with a paper tissue to remove excess water from the hydrogel surface.</li>
		<li>Weigh the hydrogel.</li>
		<li>Calculate the swelling ratio using the following formula: 
		<img alt="Equation" src="/files/ftp_upload/55813/55813eq1.jpg" /> 
		where Wd is the weight of the dried gel (step 7.1.2.) and Ws is the weight of the wet gel (step 7.1.7). Multiply with 100 to get the swelling ratio percent.</li>
	</ol>
	</li>
	<li>pH and temperature stability.
	<ol>
		<li>Transfer 5 mL of PBS to a 15-mL reaction tube.</li>
		<li>Adjust the solution to the appropriate pH (e.g., pH 2, 7, or 10) with NaOH and HCl. Bring the solution to the appropriate temperature (e.g., RT, 37 &#176;C, or 80 &#176;C).</li>
		<li>Transfer the hydrogel that is to be investigated for its stability into the appropriate solution. Readjust the pH if necessary. 
		NOTE: All hydrogels should be completely swollen at this point (see the swelling ratio) to prevent the incorrect interpretation of the weight due to the swelling of the material.</li>
		<li>After certain time intervals, remove the hydrogel from the solution (e.g., each hour for up to 2 days), dry it with a highly absorbent paper tissue to remove excess water, and weigh it.</li>
	</ol>
	</li>
	<li>Enzymatic degradation.
	<ol>
		<li>Prepare a stock enzyme solution of 300 U trypsin and pepsin, as per the manufacturer&#39;s instructions.</li>
		<li>Transfer the hydrogel (e.g., from step 1.8) to the appropriate solution. 
		NOTE: All hydrogels should be completely swollen at this point to prevent the incorrect interpretation of the weight.</li>
		<li>After certain time intervals, remove the hydrogel from the solution (e.g., each hour), dry it with a highly absorbent paper tissue to remove excess water, and weigh it.</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests.

The production of macroporous matrices can be beneficial to many different fields. It has high technical and economic potential due to the defined structure of the hydrogel and the ability to control and tune specific material properties. However, the introduction of supramolecular structural elements, such as pores or channels, to a 3D template might influence the overall properties of a material, such as the swelling ratio or the stiffness. This can result in the undesired decomposition, degradation, or breakdown of th...

Hydrogels are materials that form insoluble 3D networks capable of binding large amounts of water. Such materials can provide excellent environmental conditions for living cells. Currently, there is increasing interest in the generation of three-dimensional hydrogel structures and in the development of processes to tailor their chemical and physical properties. Once this is achieved, a template for the growth of cells and the manipulation of cellular behavior can be generated1,2,3,4. These 3D structures not only create a more natural and realistic environment than conventional two-dimensional approaches, but they also reveal new possibilities for the growth of stem cells or tumor models5. Different materials possess a range of characteristics that mainly depend upon the pore size of the gel6. The pores play a crucial role in cell culture applications, tissue engineering, and the directed growth of stem cells. For example, oxygen and nutrients diffuse through the matrix, and adequate amounts must be able to reach the cells7. On the other hand, harmful metabolites must be removed as quickly as possible, and sufficient space for cell growth must be available7. Consequently, the properties of the material, and thus the pore size, severely influence the potential benefit and possible applications of the matrix. Depending upon the properties of the material, different cell-growth processes can occur in 3D cell culture, including the formation of neuronal structures; the growth and differentiation of skin or bone cells; and the directed growth of special stem cell lines, like hepatocytes or fibroblasts2,3,8,9,10,11. Another crucial point influencing the possible application of a material is its stability towards external stimuli12. For example, the hydrogel must maintain its mechanical integrity in cell culture media or the human body.
In recent years, research on 3D cell culture hydrogels intensified, and many studies were carried out to resolve the 3D architectures of the systems13. Hydrogels composed of chemically synthesized components are most commonly investigated because they can be easily synthesized and chemically modified and they exhibit high stability (see Zhu et al., 2011 for a review)5. However, proteins have many beneficial properties: as so-called &#34;precision polymers,&#34; they are biocompatible; they have a defined length; they are relatively easy to modify; and they have a large number of target sites14,15. In this regard, highly specific, innovative structures can be generated for application in many fields. In this study, a protein-based hydrogel16 was used to demonstrate the ability of well-established methods to influence the 3D architecture of the material. Furthermore, the capability of and applicability to pore generation was also investigated.
Many different techniques are available to modify 3D structures, including both simple methods and sophisticated, highly specialized techniques from different fields of material science. A widespread technique is the use of electrospinning to generate well-defined structures17. Charged fibers are pulled from a solution by an electric field and then solidify upon exposure to oxygen. In this way, fibers in the range of several nanometers up to several microns can be produced. Additional techniques to tune the size, structure, and distribution of the pores within the matrix are soft lithography, photolithography, hydrodynamic focusing, electro-spraying, and bio-printing18,19,20. A significant drawback of these techniques is their dependency upon specific, expensive equipment and special chemicals or materials. Furthermore, experience with these techniques is often not directly transferrable to protein-based materials, and many of the chemicals and methods are not cell compatible.
On the other hand, many techniques do not rely upon special equipment, making them easier and cheaper to apply and to reproduce. A widespread method for structure manipulation is solvent casting21,22,23. Particles are added prior to the polymerization reaction and are distributed homogenously to saturate the solution. After the polymerization, a change of conditions, such as a dilution or a pH change, leads to the solvation of the particles, while the pores remain within the material. The chemicals used in these techniques, such as salt, sugar, paraffin, gelatin, and chalk, are cheap and readily available. In freeze-drying, swollen hydrogels are frozen. The subsequent sublimation of the liquid phases under a vacuum is then performed23,24,25. Water sublimation from the network is gentle enough to maintain the specific 3D structures of the material. In gas foaming, a solution is streamed with a gas while the polymerization takes place, leaving pores within the gel21. The size and distribution of the pores can be adjusted depending upon the gas stream.
To form the protein hydrogel, BSA is reacted with tetrakis (hydroxymethyl) phosphonium chloride (THPC) in a Mannich-type reaction to allow for the formation of covalent bonds between primary amines and the hydroxy groups of the four-armed linker molecule26. Possible harmful intermediates are removed by excessive washing of the material after the reaction occurs.
This study demonstrates the possibility of treating a BSA-based material with different techniques to manipulate and tailor the size of the pores. Each of the techniques can be used in any laboratory worldwide, as no special equipment is necessary. In addition, different parameters, such as swelling ratio, enzymatic degradability, pH stability, and temperature sensitivity, were examined and compared to each other, especially respect to the influence of the different techniques on the generation of 3D architectures. Finally, the materials were functionalized with cell-adhesive peptides to investigate the possible application of the materials to cell culture. Two different model cell lines were used: A549 and MCF7.

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easy manipulation of architectures in protein-based hydrogels for cell culture applications

Hydrogels are recognized as promising materials for cell culture applications due to their ability to provide highly hydrated cell environments. The field of 3D templates is rising due to the potential resemblance of those materials to the natural extracellular matrix. Protein-based hydrogels are particularly promising because they can easily be functionalized and can achieve defined structures with adjustable physicochemical properties. However, the production of macroporous 3D templates for cell culture applications using natural materials is often limited by their weaker mechanical properties compared to those of synthetic materials. Here, different methods were evaluated to produce macroporous bovine serum albumin (BSA)-based hydrogel systems, with adjustable pore sizes in the range of 10 to 70 &#181;m in radius. Furthermore, a method to generate channels in this protein-based material that are several hundred microns long was established. The different methods to produce pores, as well as the influence of pore size on material properties such as swelling ratio, pH, temperature stability, and enzymatic degradation behavior, were analyzed. Pore sizes were investigated in the native, swollen state of the hydrogels using confocal laser scanning microscopy. The feasibility for cell culture applications was evaluated using a cell-adhesive RGD peptide modification of the protein system and two model cell lines: human breast cancer cells (A549) and adenocarcinomic human alveolar basal epithelial cells (MCF7).

Hydrogel development has become one of the most prominent fields in material research-related biological studies, with thousands of entries indexed in scientific research archives. Although the behavior of many systems is well studied, the manipulation of 3D networks, especially of sensitive protein-based materials, is often a major issue in material science. Another commonly underestimated challenge is the correct measurement of the native structure of a material using cryo electron micr...

Watch this Scientific Journal Video about Easy Manipulation of Architectures in Protein-based Hydrogels for Cell Culture Applications at JoVE.com

Easy Manipulation of Architectures in Protein-based Hydrogels for Cell Culture Applications

Different methods to manipulate three-dimensional architecture in protein-based hydrogels are evaluated here with respect to material properties. The macroporous networks are functionalized with a cell-adhesive peptide, and their feasibility in cell culture is evaluated using two different model cell lines.

Hydrogels are recognized as promising materials for cell culture applications due to their ability to provide highly hydrated cell environments. The field ...

easy-manipulation-architectures-protein-based-hydrogels-for-cell

Universidad de Cartagena UDEC

Research

JoVE Journal

Biochemistry

6.7K Views.  Ulm University. Different methods to manipulate three-dimensional architecture in protein-based hydrogels are evaluated here with respect to material properties. The macroporous networks are functionalized with a cell-adhesive peptide, and their feasibility in cell culture is evaluated using two different model cell lines.

Video: Easy Manipulation of Architectures in Protein-based Hydrogels for Cell Culture Applications

Preparation of Homogeneous MALDI Samples for Quantitative Applications

This protocol demonstrates a simple sample preparation to reduce spatial heterogeneity in ion signals during matrix-assisted laser desorption/ionization (MALDI) mass spectrometry. The heterogeneity of ion signals is a severe problem in MALDI, which results in poor data reproducibility and makes MALDI unsuitable for quantitative analysis. By regulating sample plate temperature during sample preparation, thermal-induced hydrodynamic flows inside droplets of sample solution are able to reduce the heterogeneity problem. A room-temperature sample preparation chamber equipped with a temperature-regulated copper base block that holds MALDI sample plates facilitates precise control of the sample drying condition. After drying of sample droplets, the temperature of sample plates is returned to room temperature and removed from the chamber for subsequent mass spectrometric analysis. The areas of samples are examined with MALDI-imaging mass spectrometry to obtain the spatial distribution of all components in the sample. In comparison with the conventional dried-droplet method that prepares samples under ambient conditions without temperature control, the samples prepared with the method demonstrated herein show significantly better spatial distribution and signal intensity. According to observations using carbohydrate and peptide samples, decreasing substrate temperature while maintaining the surroundings at ambient temperature during the drying process can effectively reduce the heterogeneity of ion signals. This method is generally applicable to various combinations of samples and matrices.

A protocol for reducing spatial heterogeneities of ion signals in MALDI mass spectrometry by regulating substrate temperature during sample drying processes is demonstrated.

This protocol demonstrates a simple sample preparation to reduce spatial heterogeneity in ion signals during matrix-assisted laser desorption/ionization ...

Sample Preparation for Mass Spectrometry-based Identification of RNA-binding Regions

Noncoding RNAs play important roles in several nuclear processes, including regulating gene expression, chromatin structure, and DNA repair. In most cases, the action of noncoding RNAs is mediated by proteins whose functions are in turn regulated by these interactions with noncoding RNAs. Consistent with this, a growing number of proteins involved in nuclear functions have been reported to bind RNA and in a few cases the RNA-binding regions of these proteins have been mapped, often through laborious, candidate-based methods.
Here, we report a detailed protocol to perform a high-throughput, proteome-wide unbiased identification of RNA-binding proteins and their RNA-binding regions. The methodology relies on the incorporation of a photoreactive uridine analog in the cellular RNA, followed by UV-mediated protein-RNA crosslinking, and mass spectrometry analyses to reveal RNA-crosslinked peptides within the proteome. Although we describe the procedure for mouse embryonic stem cells, the protocol should be easily adapted to a variety of cultured cells.

We describe a protocol to identify RNA-binding proteins and map their RNA-binding regions in live cells using UV-mediated photocrosslinking and mass spectrometry.

Noncoding RNAs play important roles in several nuclear processes, including regulating gene expression, chromatin structure, and DNA repair. In most ...

Single-molecule Manipulation of G-quadruplexes by Magnetic Tweezers

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA processing, and telomere elongation. During these processes, various proteins bind and resolve G4 structures to perform their function. As the function of G4 often depends on the stability of its folded structure, it is important to investigate how G4 binding proteins regulate the stability of G4. This work presents a method to manipulate single G4 molecules using magnetic tweezers, which enables studies of the regulation of G4 binding proteins on a single G4 molecule in real time. In general, this method is suitable for a wide scope of applications in studies for proteins/ligands interactions and regulations on various DNA or RNA secondary structures.

A single-molecule magnetic tweezers platform to manipulate G-quadruplexes is reported, which allows for the study of G4 stability and regulation by various proteins.

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA ...

A Fluorogenic Peptide Cleavage Assay to Screen for Proteolytic Activity: Applications for coronavirus spike protein activation

Enveloped viruses such as coronaviruses or influenza virus require proteolytic cleavage of their fusion protein to be able to infect the host cell. Often viruses exhibit cell and tissue tropism and are adapted to specific cell or tissue proteases. Moreover, these viruses can introduce mutations or insertions into their genome during replication that may affect the cleavage, and thus can contribute to adaptations to a new host. Here, we present a fluorogenic peptide cleavage assay that allows a rapid screening of peptides mimicking the cleavage site of viral fusion proteins. The technique is very flexible and can be used to investigate the proteolytic activity of a single protease on many different substrates, and in addition, it also allows exploration of the activity of multiple proteases on one or more peptide substrates. In this study, we used peptides mimicking the cleavage site motifs of the coronavirus spike protein. We tested human and camel derived Middle East Respiratory Syndrome coronaviruses (MERS-CoV) to demonstrate that single and double substitutions in the cleavage site can alter the activity of furin and dramatically change cleavage efficiency. We also used this method in combination with bioinformatics to test furin cleavage activity of feline coronavirus spike proteins from different serotypes and strains. This peptide-based method is less labor- and time intensive than conventional methods used for the analysis of proteolytic activity for viruses, and results can be obtained within a single day.

We present a fluorogenic peptide cleavage assay that allows a rapid screening of the proteolytic activity of proteases on peptides representing the cleavage site of viral fusion peptides. This method can also be used on any other amino acid motif within a protein sequence to test for the protease activity.

Enveloped viruses such as coronaviruses or influenza virus require proteolytic cleavage of their fusion protein to be able to infect the host cell. Often ...

Study on the Metabolism of Six Systemic Insecticides in a Newly Established Cell Suspension Culture Derived from Tea (Camellia Sinensis L.) Leaves

A platform for studying insecticide metabolism using in vitro tissues of tea plant was developed. Leaves from sterile tea plantlets were induced to form loose callus on Murashige and Skoog (MS) basal media with the plant hormones 2,4-dichlorophenoxyacetic acid (2,4-D, 1.0 mg L-1) and kinetin (KT, 0.1 mg L-1). Callus formed after 3 or 4 rounds of subculturing, each lasting 28 days. Loose callus (about 3 g) was then inoculated into B5 liquid media containing the same plant hormones and was cultured in a shaking incubator (120 rpm) in the dark at 25 &#177; 1 &#176;C. After 3&#8722;4 subcultures, a cell suspension derived from tea leaf was established at a subculture ratio ranging between 1:1 and 1:2 (suspension mother liquid: fresh medium). Using this platform, six insecticides (5 &#181;g mL-1 each thiamethoxam, imidacloprid, acetamiprid, imidaclothiz, dimethoate, and omethoate) were added into the tea leaf-derived cell suspension culture. The metabolism of the insecticides was tracked using liquid chromatography and gas chromatography. To validate the usefulness of the tea cell suspension culture, the metabolites of thiamethoxan and dimethoate present in treated cell cultures and intact plants were compared using mass spectrometry. In treated tea cell cultures, seven metabolites of thiamethoxan and two metabolites of dimethoate were found, while in treated intact plants, only two metabolites of thiamethoxam and one of dimethoate were found. The use of a cell suspension simplified the metabolic analysis compared to the use of intact tea plants, especially for a difficult matrix such as tea.

Study on the Metabolism of Six Systemic Insecticides in a Newly Established Cell Suspension Culture Derived from Tea Camellia Sinensis L. Leaves

This work presents a protocol for establishing a cell suspension culture derived from tea (Camellia sinensis L.) leaves that can be used to study the metabolism of external compounds that can be taken up by the whole plant, such as insecticides.

A platform for studying insecticide metabolism using in vitro tissues of tea plant was developed. Leaves from sterile tea plantlets were induced to form ...

Quantitative Analysis of the Cellular Lipidome of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

Lipids are structurally diverse amphipathic molecules that are insoluble in water. Lipids are essential contributors to the organization and function of biological membranes, energy storage and production, cellular signaling, vesicular transport of proteins, organelle biogenesis, and regulated cell death. Because the budding yeast Saccharomyces cerevisiae is a unicellular eukaryote amenable to thorough molecular analyses, its use as a model organism helped uncover mechanisms linking lipid metabolism and intracellular transport to complex biological processes within eukaryotic cells. The availability of a versatile analytical method for the robust, sensitive, and accurate quantitative assessment of major classes of lipids within a yeast cell is crucial for getting deep insights into these mechanisms. Here we present a protocol to use liquid chromatography coupled with tandem mass spectrometry (LC-MS/MS) for the quantitative analysis of major cellular lipids of S. cerevisiae. The LC-MS/MS method described is versatile and robust. It enables the identification and quantification of numerous species (including different isobaric or isomeric forms) within each of the 10 lipid classes. This method is sensitive and allows identification and quantitation of some lipid species at concentrations as low as 0.2 pmol/&#181;L. The method has been successfully applied to assessing lipidomes of whole yeast cells and their purified organelles. The use of alternative mobile phase additives for electrospray ionization mass spectrometry in this method can increase the efficiency of ionization for some lipid species and can be therefore used to improve their identification and quantitation.

Quantitative Analysis of the Cellular Lipidome of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

We present a protocol using liquid chromatography coupled with tandem mass spectrometry to identify and quantify major cellular lipids in Saccharomyces cerevisiae. The described method for a quantitative assessment of major lipid classes within a yeast cell is versatile, robust, and sensitive.

Lipids are structurally diverse amphipathic molecules that are insoluble in water. Lipids are essential contributors to the organization and function of ...

Transverse Sectioning of Mature Rice (Oryza sativa L.) Kernels for Scanning Electron Microscopy Imaging Using Pipette Tips as Immobilization Support

Starch granules (SGs) exhibit different morphologies depending on the plant species, especially in the endosperm of the Poaceae family. Endosperm phenotyping can be used to classify genotypes based on SG morphotype using scanning electron microscopic (SEM) analysis. SGs can be visualized using SEM by slicing through the kernel (pericarp, aleurone layers, and endosperm) and exposing the organellar contents. Current methods require the rice kernel to be embedded in plastic resin and sectioned using a microtome or embedded in a truncated pipette tip and sectioned by hand using a razor blade. The former method requires specialized equipment and is time-consuming, while the latter introduces a new host of problems depending on rice genotype. Chalky rice varieties, particularly, pose a problem for this type of sectioning due to the friable nature of their endosperm tissue. Presented here is a technique for preparing translucent and chalky rice kernel sections for microscopy, requiring only pipette tips and a scalpel blade. Preparing the sections within the confines of a pipette tip prevents rice kernel endosperm from shattering (for translucent or &#8216;vitreous&#8217; phenotypes) and crumbling (for chalky phenotypes). Using this technique, endosperm cell patterning and the structure of intact SGs can be observed.

Transverse Sectioning of Mature Rice Oryza sativa L. Kernels for Scanning Electron Microscopy Imaging Using Pipette Tips as Immobilization Support

This protocol allows for the preparation of transverse sections of cereal seeds (e.g., rice) for the analysis of endosperm and starch granule morphology using scanning electron microscopy.

Starch granules (SGs) exhibit different morphologies depending on the plant species, especially in the endosperm of the Poaceae family. Endosperm ...

Quantitative Metabolomics of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

Metabolomics is a methodology used for the identification and quantification of many low-molecular-weight intermediates and products of metabolism within a cell, tissue, organ, biological fluid, or organism. Metabolomics traditionally focuses on water-soluble metabolites. The water-soluble metabolome is the final product of a complex cellular network that integrates various genomic, epigenomic, transcriptomic, proteomic, and environmental factors. Hence, the metabolomic analysis directly assesses the outcome of the action for all these factors in a plethora of biological processes within various organisms. One of these organisms is the budding yeast Saccharomyces cerevisiae, a unicellular eukaryote with the fully sequenced genome. Because S. cerevisiae is amenable to comprehensive molecular analyses, it is used as a model for dissecting mechanisms underlying many biological processes within the eukaryotic cell. A versatile analytical method for the robust, sensitive, and accurate quantitative assessment of the water-soluble metabolome would provide the essential methodology for dissecting these mechanisms. Here we present a protocol for the optimized conditions of metabolic activity quenching in and water-soluble metabolite extraction from S. cerevisiae cells. The protocol also describes the use of liquid chromatography coupled with tandem mass spectrometry (LC-MS/MS) for the quantitative analysis of the extracted water-soluble metabolites. The LC-MS/MS method of non-targeted metabolomics described here is versatile and robust. It enables the identification and quantification of more than 370 water-soluble metabolites with diverse structural, physical, and chemical properties, including different structural isomers and stereoisomeric forms of these metabolites. These metabolites include various energy carrier molecules, nucleotides, amino acids, monosaccharides, intermediates of glycolysis, and tricarboxylic cycle intermediates. The LC-MS/MS method of non-targeted metabolomics is sensitive and allows the identification and quantitation of some water-soluble metabolites at concentrations as low as 0.05 pmol/&#181;L. The method has been successfully used for assessing water-soluble metabolomes of wild-type and mutant yeast cells cultured under different conditions.

Quantitative Metabolomics of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

We present a protocol for the identification and quantitation of major classes of water-soluble metabolites in the yeast Saccharomyces cerevisiae. The described method is versatile, robust, and sensitive. It allows the separation of structural isomers and stereoisomeric forms of water-soluble metabolites from each other.

Metabolomics is a methodology used for the identification and quantification of many low-molecular-weight intermediates and products of metabolism within ...

Continuous Fluorescence-Based Endonuclease-Coupled DNA Methylation Assay to Screen for DNA Methyltransferase Inhibitors

DNA methylation, a form of epigenetic gene regulation, is important for normal cellular function. In cells, proteins called DNA methyltransferases (DNMTs) establish and maintain the DNA methylation pattern. Changes to the normal DNA methylation pattern are linked to cancer development and progression, making DNMTs potential cancer drug targets. Thus, identifying and characterizing novel small molecule inhibitors of these enzymes is of great importance. This paper presents a protocol that can be used to screen for DNA methyltransferase inhibitors. The continuous coupled kinetics assay allows for initial velocities of DNA methylation to be determined in the presence and absence of potential small molecule inhibitors. The assay uses the methyl-sensitive endonuclease Gla I to couple methylation of a hemimethylated DNA substrate to fluorescence generation.
This continuous assay allows for enzyme activity to be monitored in real time. Conducting the assay in small volumes in microtiter plates reduces the cost of reagents. Using this assay, a small example screen was conducted for inhibitors of DNMT1, the most abundant DNMT isozyme in humans. The highly substituted anthraquinone natural product, laccaic acid A, is a potent, DNA-competitive inhibitor of DNMT1. Here, we examine three potential small molecule inhibitors &#8212; anthraquinones or anthraquinone-like molecules with one to three substituents &#8212; at two concentrations to describe the assay protocol. Initial velocities are used to calculate the percent activity observed in the presence of each molecule. One of three compounds examined exhibits concentration-dependent inhibition of DNMT1 activity, indicating that it is a potential inhibitor of DNMT1.

DNA methyltransferases are potential cancer drug targets. Here, a protocol is presented to assess small molecules for DNA methyltransferase inhibition. This assay utilizes an endonuclease to couple DNA methylation to fluorescence generation and allows for enzyme activity to be monitored in real time.&#160;

DNA methylation, a form of epigenetic gene regulation, is important for normal cellular function. In cells, proteins called DNA methyltransferases (DNMTs) ...

A Triple Primary Cell Culture Model of the Human Blood-Brain Barrier for Studying Ischemic Stroke In Vitro

Ischemic stroke is a major cause of death and disability worldwide with limited therapeutic options. The neuropathology of ischemic stroke is characterized by an interruption in blood supply to the brain leading to cell death and cognitive dysfunction. During and after ischemic stroke, blood-brain barrier (BBB) dysfunction facilitates injury progression and contributes to poor patient recovery. Current BBB models primarily include endothelial monocultures and double co-cultures with either astrocytes or pericytes.
Such models lack the ability to fully imitate a dynamic brain microenvironment, which is essential for cell-to-cell communication. Additionally, commonly used BBB models often contain immortalized human endothelial cells or animal-derived (rodent, porcine, or bovine) cell cultures that pose translational limitations. This paper describes a novel well-insert-based BBB model containing only primary human cells (brain microvascular endothelial cells, astrocytes, and brain vascular pericytes) enabling the investigation of ischemic brain injury in vitro. 
The effects of oxygen-glucose deprivation (OGD) on barrier integrity were assessed by passive permeability, transendothelial electrical resistance (TEER) measurements,and direct visualization of hypoxic cells. The presented protocol offers a distinct advantage inmimicking the intercellular environment of the BBB in vivo, serving as a more realistic in vitro BBB model for developing new therapeutic strategies in the setting of ischemic brain injury.

A Triple Primary Cell Culture Model of the Human Blood-Brain Barrier for Studying Ischemic Stroke In Vitro

Here, we describe the method for establishing a triple cell culture model of the blood-brain barrier based on primary human brain microvascular endothelial cells, astrocytes, and pericytes. This multicellular model is suitable for studies of neurovascular unit dysfunction during ischemic stroke in vitro or for the screening of drug candidates.

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A Triple Primary Cell Culture Model of the Human Blood-Brain Barrier for Studying Ischemic Stroke In Vitro