Herstellvorschrift Inkubationspuffer, Assay-Puffer und 0,6 N TCA-Lösung als ready-to-use-Lösungen zur Verfügung gestellt. Die FITC-Casein Substrat - Es ist zu Aliquot der FITC-Casein Substrat in kleineren Mengen bei der Ankunft auf wiederholtes Einfrieren und Auftauen zu vermeiden empfohlen. Wenn die FITC-Casein Substrat wiederholt Gefrier-Auftau-Zyklen unterworfen ist, wird ein leichter Anstieg in den Hintergrund treten, dadurch Senkung der Empfindlichkeit. Die Aliquots bei -20 ° C gelagert werden und vor Licht geschützt. Jede Probe, erfordert leer oder Kontrollreaktion 20 ul der FITC-Casein Substrat. Frost / Tau-sowie kräftiges Mischen oder Schütteln kann der FITC aus dem Casein was zu einer höheren Hintergrund Lesen und verursacht weniger Substrat zur Verfügung zu stehen Proteaseschnittstelle trennen. Die Fluorescein-Isothiocyanat (FITC) Control Solution-Der FITC-Steuerung kann in Assay-Puffer auf die geeignete Konzentration aufgelöst werden. Diese Lösung sollte frisch zubereitet werden und vor Licht geschützt. Finale der Trypsin-Control-Lösung - 100 l von 1 mM HCl in das Fläschchen von Trypsin, Protease Control (Product Code T 6567). Mix kurz, um sicherzustellen, das Trypsin gelöst ist. Add 900 ul Inkubationspuffer und gut mischen. Alternativ können auch andere Puffer verwendet werden, wenn für den Test gewünscht wird. Die letzte Arbeit Konzentration der Trypsin ist 20 ug / ul. Wenn Sie bereit sind, die Trypsin-Control-Lösung vorbereiten, kombinieren einen aliquoten Teil der sauren Trypsin-Lösung mit der richtigen Menge der Inkubationspuffer (1 Teil angesäuert Trypsin zu 9 Teilen Puffer). Das Speichern der Trypsin unter sauren Bedingungen erhöht die Stabilität des Trypsin. Die Trypsin-Lösung ist temperaturempfindlich ist und nicht in niedrigen Konzentrationen stabil. Bitte überprüfen Sie die Kits technischen Merkblatt für die Lagerung und die Stabilität dieser Lösungen vorbereitet. Verfahren <ol><li> Am einfachsten ist es, die Kontrollen, Blindproben und Proben aller Vorbereitung auf einmal. Für jede Probe werden 20 ul Inkubationspuffer, 20 ul von FITC-Casein Substrat, und 10 ul der Probe zu einem Reaktionsgefäß. Für Proben mit hohen Protease-Aktivität kann Probenverdünnung erforderlich sein. </li><li> Bereiten Sie entsprechende Kontrollproben (siehe Kontrollproben) durch Zugabe von 20 ul Inkubationspuffer, 20 ul von FITC-Casein Substrat, und 10 ul der Kontrollprobe ein Mikrozentrifugenröhrchen. </li><li> Bereiten Sie eine leere Probe durch Zugabe von 20 ul Inkubationspuffer, 20 ul von FITC-Casein Substrat, und 10 ul Reinstwasser auf ein Mikrozentrifugenröhrchen. </li><li> Vorsichtig mischen jedes Rohr und bei 37 ° C im Dunkeln für 60 Minuten. Seien Sie vorsichtig, nicht zu kräftig mischen, da übermäßige Turbulenzen hohen Fluoreszenz-Hintergrund führen können und reduzieren Sie die Empfindlichkeit des Tests. Hinweis: Die Inkubationszeit kann bis zu 24 Stunden verlängert werden, um die Empfindlichkeit zu erhöhen. Achten Sie darauf, 24 Stunden überschreiten, wie die FITC-Casein kann beginnen sich zu zersetzen, was zu hohen Fluoreszenz-Hintergrund. </li><li> Nach der Inkubation add 150 ul der 0,6 N Trichloressigsäurelösung jedes Mikrozentrifugenröhrchen. TCA ist stark ätzend und sollten so behandelt, während das Tragen der entsprechenden Schutzausrüstung. </li><li> Vorsichtig mischen und bei 37 ° C im Dunkeln für 30 Minuten. </li><li> Zentrifugieren für 10 Minuten bei 10.000 x g. Der Überstand enthält die Säure löslich, FITC markierten Fragmente und ist für die Fluoreszenz-Messung verwendet. </li></ol> Fluoreszenz-Messungen Diese Methoden können skaliert werden oder nach unten entsprechend den Anforderungen der Instrumentierung zur Verfügung. Zum Vergleich mit einer Standardkurve mit den entsprechenden Kontrollproben vorbereitet, subtrahieren die Fluoreszenz Lesung der Blindprobe (FLUblank) aus dem Wert von jeder Testprobe (FLUtest). Küvetten <ol><li> Je 10 ul des Überstandes (Schritt 7) und 1 ml der Assay-Puffer in eine geeignete Küvette und vorsichtig mischen. Hinweis: Die Lösung des Überstands und Assay-Puffer können im Dunkeln bei 2-8 ° C gelagert werden bis zu 24 Stunden vor der Messung der Fluoreszenz. </li><li> Notieren Sie sich die Fluoreszenz-Intensität mit einer Anregung bei 485 nm und Überwachung der Emissions-Wellenlänge von 535 nm. </li></ol> Multiwell Platten <ol><li> Je 10 ul des Überstandes (Schritt 7) und 1 ml der Assay-Puffer in ein geeignetes Röhrchen oder Fläschchen geben und vorsichtig mischen Hinweis: Die Lösung des Überstands und Assay-Puffer können im Dunkeln bei 2-8 ° C für bis abgelegt bis 24 Stunden vor der Messung der Fluoreszenz. </li><li> Transfer-200 ul in eine Vertiefung einer schwarzen 96-Well-Platte. Notieren Sie sich die Fluoreszenz-Intensität mit einer Anregung bei 485 nm und Überwachung der Emissions-Wellenlänge von 535 nm. Oder </li></ol><ol><li> Pipette 2 ul des Überstandes (Schritt 7) und 200 ul der Assay BUffer in eine Vertiefung einer schwarzen 96-Well-Platte. Hinweis: Die Lösung des Überstands und Assay-Puffer können im Dunkeln bei 2-8 ° C gelagert werden bis zu 24 Stunden vor der Messung der Fluoreszenz. </li><li> Notieren Sie sich die Fluoreszenz-Intensität mit einer Anregung bei 485 nm und Überwachung der Emissions-Wellenlänge von 535 nm. </li></ol> Kontrollproben Die Trypsin-Control-Lösung kann verwendet werden um zu bestätigen, ist der Test korrekt durchführen, um die Nachweisgrenze zu bestimmen, oder erstellen Sie einen allgemeinen Standard-Kurve liegen. Für den Test eines anderen, spezifische Protease, ist es empfehlenswert, eine Kontroll-Lösung, welche die spezifische Protease in den entsprechenden Inkubationspuffer vorzubereiten. Die Nachweisgrenze des Tests ist die Menge der Protease, die eine deutliche Fluoreszenz Lesung über den Wert mit der Blindprobe erhalten produziert. Die Nachweisgrenze ist abhängig von der Empfindlichkeit der Instrumente. Serielle Verdünnungen der Trypsin-Control-Lösung kann verwendet werden, um die Control-Lösungen zu generieren. Eine Lesung in Höhe von 120% des Wertes der Blindprobe erhält man als signifikant betrachtet. Routinemäßig wurde eine Nachweisgrenze von 5 ng Trypsin mit diesem Verfahren erhalten. Es wird empfohlen, mindestens eine 5 ng Trypsin-Steuerung mit jedem Test ausgeführt werden. Ein Trypsin-Lösung mit einer Konzentration von 0,5 ug / ml würden in der gewünschten 5 ng Trypsin im Test führen. Ein 40-fachen Verdünnung des Trypsin-Control-Lösung (20 ug / ml) ergibt sich ein 0,5 ug / ml Kontroll-Lösung, dh ein Teil der Trypsin-Control-Lösung zu 39 Teilen Inkubationspuffer. <img src="/files/ftp_upload/1514/figure1.jpg" alt="Abbildung 1" /> Abbildung 1 (Standard Curve von Trypsin Activity): Die Trypsin-Control-Lösung (20 pg / ml) kann auch verwendet werden, um eine Standardkurve, indem serielle Verdünnungen zu erzeugen (siehe Abbildung 1). Die Fluoreszenz Lesen von jeder Kontrolle Probe wurde durch Subtraktion der Fluoreszenz-Lesung der Blindprobe (FLUblank) aus dem Wert der einzelnen Kontrollprobe (FLUcontrol) korrigiert. Typische Standardkurve für Trypsin (Product Code T 6567) mit diesem Kit und Kontrollproben von 0,15 pg / ml (1,5 ng) auf 2,5 pg / ml (25 ng). Diese Kurve wurde gemäß dem beschriebenen Verfahren, mit einer Inkubationszeit von 30 Minuten für Schritt 4 erzeugt. Fluoreszenz-Messungen wurden in einer Multi-Well-Platte. Fluoresceinisothiocyanat ist als Kontrolle für mögliche Gerätekalibrierung (siehe entsprechende Anweisungen des Herstellers) oder die Bestimmung der Linearitätsbereich der FITC-Signal zur Verfügung gestellt.

<ol>
	<li>Twining, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescein+Isothiocyanate-Labeled+Casein+Assay+for+Proteolytic+Enzymes.">Fluorescein Isothiocyanate-Labeled Casein Assay for Proteolytic Enzymes.</a> Anal. Biochem. 143, 30-34 (1984).</li></ol>

Der Autor ist Mitarbeiter von Sigma Aldrich, dass Reagenzien und Instrumente in diesem Artikel verwendet produziert.

Wir haben gerade ein Beispiel Assay gezeigt mit der Protease Fluorescent Detection Kit für Protease-Aktivität in einer biologischen Probe zu erkennen. Dieses Kit wurde optimiert, um ein breites Spektrum von Proteasen in physiologischen Anwendungen gefunden zu erkennen. Es eignet sich zum Nachweis von Serin, Cystein-, Metallo-und Aspartylproteasen, aber Modifikationen erforderlich, um einige spezifische Proteasen zu erkennen.

<table border="1">
<tr>
<td>Material Name</td>
<td>Type</td>
<td>Company</td>
<td>Catalogue Number</td>
<td>Comments (optional)</td>
</tr>
<tr>
<td><a href="http://www.sigmaaldrich.com/catalog/ProductDetail.do?lang=en&N4=PF0100|SIGMA&N5=SEARCH_CONCAT_PNO|BRAND_KEY&F=SPEC&cm_sp=affiliate-_-Jove-_-12346">Protease Fluorescent Detection Kit</a></td>
<td>&nbsp;</td>
<td>Sigma Aldrich</td>
<td>PF0100</td>
<td>&nbsp;</td>
<tr>
<td><a href="http://www.sigmaaldrich.com/catalog/ProductDetail.do?lang=en&N4=H1758|SIGMA&N5=SEARCH_CONCAT_PNO|BRAND_KEY&F=SPEC&cm_sp=affiliate-_-Jove-_-12347">Hydrochloric Acid</a></td>
<td>&nbsp;</td>
<td>Sigma Aldrich</td>
<td>H1758</td>
<td>&nbsp;</td>
</table>

use of the protease fluorescent detection kit to determine protease activity

Die Protease Fluorescent Detection Kit bietet ready-to-use Reagenzien für den Nachweis von Protease-Aktivität. Dieser einfache Test auf Protease-Aktivität zu erkennen nutzt Kasein mit Fluoresceinisothiocyanat (FITC) als Substrat bezeichnet. Protease-Aktivität führt zur Spaltung der FITC-markiertem Casein Substrat in kleinere Fragmente, die nicht ausfallen müssen unter sauren Bedingungen. Nach der Inkubation der Protease Probe und Substrat wird die Reaktion mit der Zugabe von Trichloressigsäure (TCA) angesäuert. Die Mischung wird dann mit dem unverdauten Substrat und bildet eine Pellet-und die kleineren, säurelöslichen Fragmenten in Lösung bleibt zentrifugiert. Der Überstand wird neutralisiert und die Fluoreszenz des FITC-markierten Fragmente gemessen wird. Die beschriebenen Kit Verfahren erkennt die Trypsin-Protease-Steuerung in einer Konzentration von etwa 0,5 pg / ml (5 ng Trypsin hinzugefügt, um die Assay). Diese Empfindlichkeit kann mit einer längeren Inkubationszeit erhöht werden, bis zu 24 Stunden. Der Test wird in Mikrozentrifugenröhrchen durchgeführt und die Verfahren für die Fluoreszenzdetektion entweder mit Küvetten oder Mikrotiterplatten zur Verfügung gestellt.

Watch this Scientific Journal Video about Use of the Protease Fluorescent Detection Kit to Determine Protease Activity at JoVE.com

Use of the Protease Fluorescent Detection Kit to Determine Protease Activity

Die Protease Fluorescent Detection Kit ist für die Messung der Protease-Aktivität mit Fluorometrie konzipiert. Es eignet sich auch zum Nachweis von Spuren von Protease-Kontamination. Die Methode basiert auf der proteolytischen hydroysis von einer proprietären Rezeptur eines FITC-markiertem Casein Substrat.

Die Nutzung der Protease Fluorescent Detection Kit für Protease-Aktivität zu bestimmen

The Protease Fluorescent Detection Kit provides ready-to-use reagents for detecting the presence of protease activity. This simple assay to detect ...

use-protease-fluorescent-detection-kit-to-determine-protease

Research

JoVE Journal

Biology

28.0K Aufrufe.  Sigma Aldrich. Die Protease Fluorescent Detection Kit ist für die Messung der Protease-Aktivität mit Fluorometrie konzipiert. Es eignet sich auch zum Nachweis von Spuren von Protease-Kontamination. Die Methode basiert auf der proteolytischen hydroysis von einer proprietären Rezeptur eines FITC-markiertem Casein Substrat.

Serie: Use of the Protease Fluorescent Detection Kit to Determine Protease Activity

Purifying Plasmid DNA from Bacterial Colonies Using the Qiagen Miniprep Kit

Plasmid DNA purification from E. coli is a core technique for molecular cloning. Small scale purification (miniprep) from less than 5 ml of bacterial culture is a quick way for clone verification or DNA isolation, followed by further enzymatic reactions (polymerase chain reaction and restriction enzyme digestion). Here, we video-recorded the general procedures of miniprep through the QIAGEN's QIAprep 8 Miniprep Kit, aiming to introducing this highly efficient technique to the general beginners for molecular biology techniques. The whole procedure is based on alkaline lysis of E. coli cells followed by adsorption of DNA onto silica in the presence of high salt. It consists of three steps: 1) preparation and clearing of a bacterial lysate, 2) adsorption of DNA onto the QIAprep membrane, 3) washing and elution of plasmid DNA. All steps are performed without the use of phenol, chloroform, CsCl, ethidium bromide, and without alcohol precipitation. It usually takes less than 2 hours to finish the entire procedure.

Purifying Plasmid-DNA aus Bakterien-Kolonien mit dem Qiagen Miniprep Kit

Plasmid DNA purification from E. coli is a core technique for molecular cloning. Small scale purification (miniprep) from less than 5 ml of bacterial ...

Forschung

Biologie

Labeling Stem Cells with Fluorescent Dyes for non-invasive Detection with Optical Imaging

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo labeling of stem cells with a fluorescent dye, subsequent intravenous injection of the labeled cells and visualization of their accumulation in specific target organs or pathologies. The presented technique demonstrates how we label human mesenchymal stem cells (hMSC) by simple incubation with the lipophilic fluorescent dye DiD (C67H103CIN2O3S) and how we label human embryonic stem cells (hESC) with the FDA approved fluorescent dye Indocyanine Green (ICG). The uptake mechanism is via adherence and diffusion of the lypophilic dye across the phospholipid cell membrane bilayer. The labeling efficiency is usually improved if the cells are incubated with the dye in serum-free media as opposed to incubation in serum-containing media. Furthermore, the addition of the transfection agent Protamine Sulfate significantly improves contrast agent uptake.

This video shows techniques for labeling of human embryonic stem cells and mesenchymal stem cells with fluorescent dyes. This technique can be used for an in vivo tracking of transplanted stem cells with optical imaging and for histopathological correlations with fluorescence microscopy.

Labeling Stammzellen mit fluoreszierenden Farbstoffen zur nicht-invasiven Detektion mit Optical Imaging

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo ...

Sigma's Non-specific Protease Activity Assay - Casein as a Substrate

Proteases break peptide bonds. In the lab, it is often necessary to measure and/or compare the activity of proteases. Sigma's non-specific protease activity assay may be used as a standardized procedure to determine the activity of proteases, which is what we do during our quality control procedures. In this assay, casein acts as a substrate. When the protease we are testing digests casein, the amino acid tyrosine is liberated along with other amino acids and peptide fragments. Folin and Ciocalteus Phenol, or Folin's reagent primarily reacts with free tyrosine to produce a blue colored chromophore, which is quantifiable and measured as an absorbance value on the spectrophotometer. The more tyrosine that is released from casein, the more the chromophores are generated and the stronger the activity of the protease. Absorbance values generated by the activity of the protease are compared to a standard curve, which is generated by reacting known quantities of tyrosine with the F-C reagent to correlate changes in absorbance with the amount of tyrosine in micromoles. From the standard curve the activity of protease samples can be determined in terms of Units, which is the amount in micromoles of tyrosine equivalents released from casein per minute.
To view this article in Chinese, <a href="http://jove.com/index/details.stp?ID=910">click here</a>

Sigma&#039;s Non-specific Protease Activity Assay - Casein as a Substrate

Proteases break peptide bonds. In the lab, it is often necessary to measure and/or compare the activity of proteases. Sigma's non-specific protease activity assay may be used as a standardized procedure to determine the activity of proteases.

Sigma Unspezifische Protease Activity Assay - Casein als Substrat

Proteases break peptide bonds. In the lab, it is often necessary to measure and/or compare the activity of proteases.  Sigma's non-specific protease ...

Direct Detection of the Acetate-forming Activity  of the Enzyme Acetate Kinase

Acetate kinase, a member of the acetate and sugar kinase-Hsp70-actin (ASKHA) enzyme superfamily1-5, is responsible for the reversible phosphorylation of acetate to acetyl phosphate utilizing ATP as a substrate. Acetate kinases are ubiquitous in the Bacteria, found in one genus of Archaea, and are also present in microbes of the Eukarya6. The most well characterized acetate kinase is that from the methane-producing archaeon Methanosarcina thermophila7-14. An acetate kinase which can only utilize PPi but not ATP in the acetyl phosphate-forming direction has been isolated from Entamoeba histolytica, the causative agent of amoebic dysentery, and has thus far only been found in this genus15,16.
In the direction of acetyl phosphate formation, acetate kinase activity is typically measured using the hydroxamate assay, first described by Lipmann17-20, a coupled assay in which conversion of ATP to ADP is coupled to oxidation of NADH to NAD+ by the enzymes pyruvate kinase and lactate dehydrogenase21,22, or an assay measuring release of inorganic phosphate after reaction of the acetyl phosphate product with hydroxylamine23. Activity in the opposite, acetate-forming direction is measured by coupling ATP formation from ADP to the reduction of NADP+ to NADPH by the enzymes hexokinase and glucose 6-phosphate dehydrogenase24.
Here we describe a method for the detection of acetate kinase activity in the direction of acetate formation that does not require coupling enzymes, but is instead based on direct determination of acetyl phosphate consumption. After the enzymatic reaction, remaining acetyl phosphate is converted to a ferric hydroxamate complex that can be measured spectrophotometrically, as for the hydroxamate assay. Thus, unlike the standard coupled assay for this direction that is dependent on the production of ATP from ADP, this direct assay can be used for acetate kinases that produce ATP or PPi.

A method for the determination of acetate kinase activity is described. This assay utilizes a direct reaction for determining enzyme activity and kinetics of acetate kinase in the acetate-forming direction with different phosphoryl acceptors. Furthermore, this method can be utilized for assaying other acetyl phosphate or acetyl-CoA utilizing enzymes.

Direkter Nachweis der Acetat-bildenden Aktivität des Enzyms Acetat-Kinase

Acetate kinase, a member of the acetate and sugar kinase-Hsp70-actin (ASKHA) enzyme superfamily1-5, is responsible for the reversible phosphorylation of ...

Protease- and Acid-catalyzed Labeling Workflows Employing 18O-enriched Water

Stable isotopes are essential tools in biological mass spectrometry. Historically, 18O-stable isotopes have been extensively used to study the catalytic mechanisms of proteolytic enzymes1-3. With the advent of mass spectrometry-based proteomics, the enzymatically-catalyzed incorporation of 18O-atoms from stable isotopically enriched water has become a popular method to quantitatively compare protein expression levels (reviewed by Fenselau and Yao4, Miyagi and Rao5 and Ye et al.6). 18O-labeling constitutes a simple and low-cost alternative to chemical (e.g. iTRAQ, ICAT) and metabolic (e.g. SILAC) labeling techniques7. Depending on the protease utilized, 18O-labeling can result in the incorporation of up to two 18O-atoms in the C-terminal carboxyl group of the cleavage product3. The labeling reaction can be subdivided into two independent processes, the peptide bond cleavage and the carboxyl oxygen exchange reaction8. In our PALeO (protease-assisted labeling employing 18O-enriched water) adaptation of enzymatic 18O-labeling, we utilized 50% 18O-enriched water to yield distinctive isotope signatures. In combination with high-resolution matrix-assisted laser desorption ionization time-of-flight tandem mass spectrometry (MALDI-TOF/TOF MS/MS), the characteristic isotope envelopes can be used to identify cleavage products with a high level of specificity. We previously have used the PALeO-methodology to detect and characterize endogenous proteases9 and monitor proteolytic reactions10-11. Since PALeO encodes the very essence of the proteolytic cleavage reaction, the experimental setup is simple and biochemical enrichment steps of cleavage products can be circumvented. The PALeO-method can easily be extended to (i) time course experiments that monitor the dynamics of proteolytic cleavage reactions and (ii) the analysis of proteolysis in complex biological samples that represent physiological conditions. PALeO-TimeCourse experiments help identifying rate-limiting processing steps and reaction intermediates in complex proteolytic pathway reactions. Furthermore, the PALeO-reaction allows us to identify proteolytic enzymes such as the serine protease trypsin that is capable to rebind its cleavage products and catalyze the incorporation of a second 18O-atom. Such "double-labeling" enzymes can be used for postdigestion 18O-labeling, in which peptides are exclusively labeled by the carboxyl oxygen exchange reaction. Our third strategy extends labeling employing 18O-enriched water beyond enzymes and uses acidic pH conditions to introduce 18O-stable isotope signatures into peptides.

Protease- and Acid-catalyzed Labeling Workflows Employing 18O-enriched Water

Stable isotope labeling workflows employing 18O-enriched water (LeO-workflows) are versatile tools for quantitative and qualitative proteomics studies. In protease-assisted (PALeO) workflows, 18O-atoms are introduced by proteolytic cleavage and carboxyl oxygen exchange reactions mediated by proteases. In the acid-catalyzed (ALeO) workflow, 18O-atoms are introduced by carboxyl oxygen exchange at low pH.

Protease-und Acid-katalysierten Labeling Workflows beschäftigt<sup&gt; 18</sup&gt; O-angereichertem Wasser

Stable isotopes are essential tools in biological mass spectrometry. Historically, 18O-stable isotopes have been extensively used to study the catalytic ...

Mouse Islet of Langerhans Isolation using a Combination of Purified Collagenase and Neutral Protease

The interrogation of beta cell gene expression and function in vitro has squarely shifted over the years from the study of rodent tumorigenic cell lines to the study of isolated rodent islets. Primary islets offer the distinct advantage that they more faithfully reflect the biology of intracellular signaling pathways and secretory responses. Whereas the method of islet isolation using tissue dissociating enzyme (TDE) preparations has been well established in many laboratories1-4, variations in the consistency of islet yield and quality from any given rodent strain limit the extent and feasibility of primary islet studies. These variations often occur as a result of the crude partially purified TDEs used in the islet isolation procedure; TDEs frequently exhibit lot-to-lot variations in activity and often require adjustments to the dose of enzyme used. A small number of reports have used purified TDEs for rodent cell isolations5, 6, but the practice is not widespread despite the routine use and advantages of purified TDEs for human islet isolations. In collaboration with VitaCyte, LLC (Indianapolis, IN), we developed a modified mouse islet isolation protocol based on that described by Gotoh7, 8, in which the TDEs are perfused directly into the pancreatic duct of mice, followed by crude tissue fractionation through a Histopaque gradient9, and isolation of purified islets. A significant difference in our protocol is the use of purified collagenase (CIzyme MA) and neutral protease (CIzyme BP) combination. The collagenase was characterized by the use of a6 fluorescence collagen degrading activity (CDA) assay that utilized fluorescently labeled soluble calf skin fibrils as substrate6. This substrate is more predictive of the kinetics of collagen degradation in the tissue matrix because it relies on native collagen as the substrate. The protease was characterized with a sensitive fluorescent kinetic assay10. Utilizing these improved assays along with more traditional biochemical analysis enable the TDE to be manufactured more consistently, leading to improved performance consistency between lots. The protocol described in here was optimized for maximal islet yield and optimal islet morphology using C57BL/6 mice. During the development of this protocol, several combinations of collagenase and neutral proteases were evaluated at different concentrations, and the final ratio of collagenase:neutral protease of 35:10 represents enzyme performance comparable to Sigma Type XI. Because significant variability in average islet yields from different strains of rats and mice have been reported, additional modifications of the TDE composition should be made to improve the yield and quality of islets recovered from different species and strains.

A detailed description of mouse islet isolation is described using the technique of in situ pancreatic ductal cannulation and perfusion of a combination of purified collagenase and neutral protease.

Maus Islet der Langerhans Isolation durch eine Kombination von gereinigtem Collagenase und Neutral Protease

The interrogation of beta cell gene expression and function in vitro has squarely shifted over the years from the study of rodent tumorigenic cell lines ...

Determining Membrane Protein Topology Using Fluorescence Protease Protection (FPP)

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established for many membrane proteins. A simple technique called fluorescence protease protection (FPP) is presented, which permits the determination of membrane protein topology in living cells. This technique has numerous advantages over other methods for determining protein topology, in that it does not require the availability of multiple antibodies against various domains of the membrane protein, does not require large amounts of protein, and can be performed on living cells. The FPP method employs the spatially confined actions of proteases on the degradation of green fluorescent protein (GFP) tagged membrane proteins to determine their membrane topology and orientation. This simple approach is applicable to a wide variety of cell types, and can be used to determine membrane protein orientation in various subcellular organelles such as the mitochondria, Golgi, endoplasmic reticulum and components of the endosomal/recycling system. Membrane proteins, tagged on either the N-termini or C-termini with a GFP fusion, are expressed in a cell of interest, which is subject to selective permeabilization using the detergent digitonin. Digitonin has the ability to permeabilize the plasma membrane, while leaving intracellular organelles intact. GFP moieties exposed to the cytosol can be selectively degraded through the application of protease, whereas GFP moieties present in the lumen of organelles are protected from the protease and remain intact. The FPP assay is straightforward, and results can be obtained rapidly.

Determining Membrane Protein Topology Using Fluorescence Protease Protection FPP

Here, we present a protocol to determine the orientation and topology of integral membrane proteins in living cells. This simple protocol relies on selective protease sensitivity of chimeras between the protein of interest and GFP.

Bestimmen Membrane Protein Topologie Mit Fluorescence Protease Schutz (FPP)

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established ...

The Use of Ex Vivo Whole-organ Imaging and Quantitative Tissue Histology to Determine the Bio-distribution of Fluorescently Labeled Molecules

Fluorescent labeling is a well-established process for examining the fate of labeled molecules under a variety of experimental conditions both in vitro and in vivo. Fluorescent probes are particularly useful in determining the bio-distribution of administered large molecules, where the addition of a small-molecule fluorescent label is unlikely to affect the kinetics or bio-distribution of the compound. A variety of methods exist to examine bio-distribution that vary significantly in the amount of effort required and whether the resulting measurements are fully quantitative, but using multiple methods in conjunction can provide a rapid and effective system for analyzing bio-distributions.
Ex vivo whole-organ imaging is a method that can be used to quickly compare the relative concentrations of fluorescent molecules within tissues and between multiple types of tissues or treatment groups. Using an imaging platform designed for live-animal or whole-organ imaging, fluorescence within intact tissues can be determined without further processing, saving time and labor while providing an accurate picture of the overall bio-distribution. This process is ideal in experiments attempting to determine the tissue specificity of a compound or for the comparison of multiple different compounds. Quantitative tissue histology on the other hand requires extensive further processing of tissues in order to create a quantitative measure of the labeled compounds. To accurately assess bio-distribution, all tissues of interest must be sliced, scanned, and analyzed relative to standard curves in order to make comparisons between tissues or groups. Quantitative tissue histology is the gold standard for determining absolute compound concentrations within tissues.
Here, we describe how both methods can be used together effectively to assess the ability of different administration methods and compound modifications to target and deliver fluorescently labeled molecules to the central nervous system1.

The Use of Ex Vivo Whole-organ Imaging and Quantitative Tissue Histology to Determine the Bio-distribution of Fluorescently Labeled Molecules

Ex vivo whole-organ imaging is a rapid method for determining the relative concentrations of fluorescently labeled compounds within and between tissues or treatment groups. Conversely, quantitative fluorescence histology, while more labor intensive, allows for the quantification of the absolute tissue levels of labeled molecules.

Die Verwendung von<em&gt; Ex Vivo</em&gt; Whole-Orgel Imaging und Quantitative Tissue Histologie des Bio-Verteilung von fluoreszenzmarkierten Moleküle zu bestimmen

Fluorescent labeling is a well-established process for examining the fate of labeled molecules under a variety of experimental conditions both in vitro ...

The Lactate Dehydrogenase Sequestration Assay &#8212; A Simple and Reliable Method to Determine Bulk Autophagic Sequestration Activity in Mammalian Cells

Bulk autophagy is characterized by the sequestration of large portions of cytoplasm into double/multi-membrane structures termed autophagosomes. Here a simple protocol to monitor this process is described. Moreover, typical results and experimental validation of the method under autophagy-inducing conditions in various types of cultured mammalian cells are provided. During bulk autophagy, autophagosomes sequester cytosol, and thereby also soluble cytosolic proteins, alongside other autophagic cargo. LDH is a stable and highly abundant, soluble cytosolic enzyme that is non-selectively sequestered into autophagosomes. The amount of LDH sequestration therefore reflects the amount of bulk autophagic sequestration. To efficiently and accurately determine LDH sequestration in cells, we employ an electrodisruption-based fractionation protocol that effectively separates sedimentable from cytosolic LDH, followed by measurement of enzymatic activity in sedimentable fractions versus whole-cell samples. Autophagic sequestration is determined by subtracting the proportion of sedimentable LDH in untreated cells from that in treated cells. The advantage of the LDH sequestration assay is that it gives a quantitative measure of the autophagic sequestration of endogenous cargo, as opposed to other methods that either involve ectopic expression of sequestration probes or semi-quantitative protease protection analyses of autophagy markers or receptors.

The Lactate Dehydrogenase Sequestration Assay &amp;#8212; A Simple and Reliable Method to Determine Bulk Autophagic Sequestration Activity in Mammalian Cells

Here a simple and well-validated protocol for measuring bulk autophagic sequestration activity in mammalian cells is described. The method is based on quantifying the proportion of lactate dehydrogenase (LDH) in sedimentable cell fractions compared to total cellular LDH levels.

Laktat-Dehydrogenase Sequestrierung Assay – Eine einfache und zuverlässige Methode zur Bestimmung Bulk Selbstverdauende Sequestration Aktivität in Säugerzellen

Bulk autophagy is characterized by the sequestration of large portions of cytoplasm into double/multi-membrane structures termed autophagosomes. Here a ...

Multiplexed Fluorescent Immunohistochemical Staining of Four Endometrial Immune Cell Types in Recurrent Miscarriage

Immunohistochemistry is the most commonly used method for the identification and visualization of tissue antigens in biological research and clinical diagnostics. It can be used to characterize various biological processes or pathologies, such as wound-healing, immune response, tissue rejection, and tissue-biomaterial interactions. However, the visualization and quantification of multiple antigens (especially for immune cells) in a single tissue section using conventional immunohistochemical (IHC) staining remains unsatisfactory. Hence, multiplexed technologies were introduced in recent years to identify multiple biological markers in a single tissue sample or an ensemble of different tissue samples. 
These technologies can be especially useful in differentiating the changes in immune cell-to-cell interactions within the endometrium between fertile women and women with recurrent miscarriages during implantation. This paper describes a detailed protocol for multiplexed fluorescence IHC staining to investigate the density and clustering of four major immune cell types simultaneously in precisely timed endometrial specimens during embryo implantation. The method includes sample preparation, multiplex optimization with markers for immune cell subtypes, and the scanning of the slides, followed by data analysis, with specific reference to detecting endometrial immune cells. 
Using this method, the density and clustering of four major immune cell types in the endometrium can be simultaneously analyzed in a single tissue section. In addition, this paper will discuss the critical factors and troubleshooting to overcome possible fluorophore interference between the fluorescent probes being applied. Importantly, the results from this multiplex staining technique can help provide an in-depth understanding of the immunologic interaction and regulation during embryo implantation.

Despite the advancements in multiplex immunohistochemistry and multispectral imaging, characterizing the density and clustering of major immune cells simultaneously in the endometrium remains a challenge. This paper describes a detailed multiplex staining protocol and imaging for the simultaneous localization of four immune cell types in the endometrium.

Gemultiplexte fluoreszierende immunhistochemische Färbung von vier endometrialen Immunzelltypen bei wiederkehrenden Fehlgeburten

Immunohistochemistry is the most commonly used method for the identification and visualization of tissue antigens in biological research and clinical ...