준비 지침 부화 버퍼, 버퍼 분석 및 0.6 N TCA 솔루션은 즉시 사용 솔루션으로 제공합니다. FITC - 카제인 기판은 - 그것은 반복 동결 - 해동주기를 피하기 위해 도착시 작은 볼륨으로 나누어지는 FITC - 카제인 기판을하는 것이 좋습니다. FITC - 카제인 기판이 반복 동결 - 해동주기를 받게되면, 백그라운드에서 약간 증가 감도를 낮추함으로써, 발생합니다. aliquots는 -20 ° C에 보관하고 빛으로부터 보호되어야합니다. 각 예제는, 공백, 또는 제어 반응은 FITC - 카제인 기판 20 μl가 필요합니다. 해동뿐만 아니라 활발한 혼합 또는 흔들림하면 FITC는 읽기와 더 적은 기판은 프로 테아제의 절단에 사용할 수로 인해 높은 배경에 발생하는 카제인에서 분리가 발생할 수 있습니다 / 프리즈. 플루오레신 isothiocyanate (FITC) 제어 솔루션 FITC 컨트롤을 적절한 농도로 분석 버퍼에 재구성 수 있습니다. 이 솔루션은 신선했고 빛으로부터 보호되어야합니다. 최종 트립신 제어 솔루션 - 트립신, 프로 테아제 제어 (제품 코드 T 6567)의 약병 1 MM HCL 100 μl를 추가합니다. 트립신이 해소되도록 간략하게 섞는다. 부화 버퍼 900 μl를 추가하고 잘 섞는다. 분석을 위해 필요한 경우 또는, 다른 버퍼를 사용할 수 있습니다. 트립신의 마지막 작업 농도는 20 μg / μl입니다. 때 트립신 제어 솔루션을 준비하는 준비 부화 버퍼 (9 부품 버퍼 1 부분 산성의 트립신)의 정확한 금액과 산성 트립신 솔루션의 나누어지는을 결합. 산성 조건 하에서 트립신을 저장하면 트립신의 안정성을 증가시킵니다. 트립신 제어 솔루션은 온도에 민감한하고 낮은 농도에서 안정되지 않습니다. 스토리지 및 이러한 준비 솔루션의 안정성을위한 키트의 기술 게시판을 참조하시기 바랍니다. 절차 <ol><li> 한 번에 모든 컨트롤, 공백, 그리고 샘플을 준비하는 가장 쉬운 그것을 것입니다. 각 테스트 샘플의 경우, microcentrifuge 관을 배양 버퍼 20 μl, FITC - 카제인 기판 20 μl 및 테스트 샘플의 10 μl를 추가합니다. 높은 단백 ​​분해 효소 활동과 테스트 샘플, 샘플 희석이 필요할 수 있습니다. </li><li> microcentrifuge 관을 배양 버퍼, FITC - 카제인 기판 20 μl 및 제어 샘플 10 μl 20 μl를 추가하여 적절한 컨트롤 샘플 (컨트롤 샘플을 참조) 준비합니다. </li><li> microcentrifuge 관을 배양 버퍼, FITC - 카제인 기판 20 μl, 그리고 ultrapure 물 10 μl 20 μl를 추가하여 빈 샘플을 준비합니다. </li><li> 부드럽게 37 60 분 어둠 속에 ° C에서 각각의 튜브와 부화를 섞는다. 과도한 난류가 높은 형광 배경을 일으킬 수 있으므로, 너무 적극적으로 믹스 및 분석의 감도를 감소하지 않도록주의하십시오. 참고 : 배양 시간이 감도를 높이기 위해 최대 24 시간까지 연장할 수 있습니다. 높은 형광 배경에 이르는, FITC - 카제인이 저하될 시작 수 있으므로 24 시간 초과하지 않도록주의하십시오. </li><li> 부화 후 각 microcentrifuge 관에 0.6 N Trichloroacetic 산성 솔루션 150 μl를 추가합니다. TCA는 매우 부식성이고 적절한 보호 장비를 착용하는 동안 처리한다. </li><li> 부드럽게 37 믹스와 부화 ° 30 분 동안 어둠 속에 C. </li><li> 10,000 X G. 10 분 동안 튜브를 원심 분리기 뜨는은 염산 가용성, FITC 레이블 조각을 포함하고 형광 측정을 위해 사용됩니다. </li></ol> 형광 측정 이 방법은 최대 크기를 조절하거나 아래로 사용할 수있는 장비의 요구 사항에 따라 수 있습니다. 해당 컨트롤 샘플 준비 표준 곡선을 비교, 각 테스트 샘플의 값 (FLUtest)에서 빈 샘플 (FLUblank)의 형광 읽는 뺍니다. Cuvettes <ol><li> 피펫 10 뜨는 (7 단계)와 적당한 쿠베트로 분석 완충액 1 ML의 μl 부드럽게 섞는다. 참고 : 최대 24 시간이 형광을 측정하기 전에를위한 뜨는 및 어세이 버퍼의 솔루션은 2-8 ° C에서 어둠에 저장되어있을 수 있습니다. </li><li> 485 NM에서 여기로 형광 강도를 기록하고 535 nm의의 방출 파장을 모니터링. </li></ol> Multiwell 플레이트 <ol><li> 뜨는 (7 단계)와 적합한 튜브 또는 유리병에 분석 완충액 1 ML 부드럽게 혼합 참고 피펫 10 μl : 뜨는 및 어세이 버퍼의 솔루션은 최대 2-8 ° C에서 어둠에 저장되어있을 수 있습니다 형광을 측정하기 전에 24 시간. </li><li> 블랙 96 잘 접시의 잘 200 μl를 전송합니다. 485 NM에서 여기로 형광 강도를 기록하고 535 nm의의 방출 파장을 모니터링. 또는 </li></ol><ol><li> 피펫 뜨는 2 μl (7 단계) 및 분석 B 200 μl블랙 96 잘 접시의 우물에 uffer. 참고 : 최대 24 시간이 형광을 측정하기 전에를위한 뜨는 및 어세이 버퍼의 솔루션은 2-8 ° C에서 어둠에 저장되어있을 수 있습니다. </li><li> 485 NM에서 여기로 형광 강도를 기록하고 535 nm의의 방출 파장을 모니터링. </li></ol> 컨트롤 샘플 트립신 제어 솔루션은 분석은 검출 한계를 결정하거나, 일반적인 표준 곡선을 만들려면 제대로 실적을 확인하는 데 사용할 수 있습니다. 다른 특정 프로 테아제의 분석을 위해, 그것은 적절한 배양 버퍼에 특정 프로 테아제를 포함하는 제어 솔루션을 준비하는 것이 좋습니다. 분석의 검출 한계는 빈 샘플과 획득의 가치 위에 상당한 형광 독서를 생산 테아제의 금액입니다. 검색의 제한은 장비의 감도에 따라 달라집니다. 트립신 제어 솔루션의 직렬 dilutions는 컨트롤 솔루션을 생성하는 데 사용할 수 있습니다. 빈 샘플 얻은 값을 120 %로 읽는 동일가 중요한 것으로 간주됩니다. 일상적으로, 트립신 5 NG의 감지 한계는이 절차를 얻은 것입니다. 이것은 하나 이상의 다섯 NG 트립신 컨트롤이 각각의 분석으로 실행하는 것이 좋습니다. 0.5 μg / ML의 농도와 트립신 제어 솔루션은 분석에서 트립신의 원하는 5 NG가 발생할 것입니다. 트립신 제어 솔루션 (20 μg / ML) 0.5 μg / ML 제어 솔루션, 인큐베이션 버퍼 39 부분 트립신 제어 솔루션 즉, 한 부분의 결과를 40 배 희석. <img src="/files/ftp_upload/1514/figure1.jpg" alt="그림 1" /> 그림 1 (트립신 활동의 표준 곡선) : 트립신 제어 솔루션 (20 μg / ML)도 (그림 1 참조) 시리얼 dilutions함으로써 표준 곡선을 생성하는 데 사용할 수 있습니다. 각 컨트롤 샘플의 형광 읽기는 각 컨트롤 샘플 (FLUcontrol)의 가치에서 빈 샘플의 형광 읽기 (FLUblank)를 빼서에 의해 수정되었습니다. 이 키트 및 0.15 μg / ML (1.5 NG)에서 2.5 μg / ML (25 NG)에 이르기까지 컨트롤 샘플을 사용하여 트립신 (제품 번호 T 6567)에 대한 전형적인 표준 곡선. 이 곡선은 4 단계 30 분 배양 시간, 설명 절차에 따라 생성되었습니다. 형광 측정은 multiwell 접시에서 만들어졌다. 플루오레신 isothiocyanate가 가능한 계측기 교정에 대한 제어 (해당 제조 업체의 지침을 참조) 또는 FITC 신호의 선형성 범위의 결정으로 제공됩니다.

<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>

저자는이 문서에서 사용하는 시약 및 도구를 만들어 시그마 알드리치의 직원입니다.

우리는 단지 생물 학적 샘플의 단백 분해 효소의 활동을 감지하는 프로 테아제 형광 탐지 키트를 사용하여 예제 분석을 보여주었다. 이 키트는 생리 애플 리케이션에서 발견 프로 테아제의 다양한 범위를 감지하기 위해 최적화되었습니다. 그것은 세린, 시스테인, 금속 및 aspartic 프로 테아제의 검출에 적합하지만, 수정은 일부 특정 프로 테아제를 감지해야 할 수 있습니다.

<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

단백 분해 효소 형광 탐지 키트를 제공 즉시 사용 테아제 활동의 존재를 검출 시약합니다. 단백 분해 효소의 활동을 감지하는이 간단한 분석은 카제인 기판으로 플루오레신 isothiocyanate (FITC)로 표시합니다. 단백 분해 효소 활동 산성 조건 하에서 침전하지 작은 조각,로 FITC - 카제인 표시 기판의 절단을 초래합니다. 단백 분해 효소 샘플 및 기판의 부화 후, 반응은 trichloroacetic 산 (TCA)의 추가와 함께 산성이다. 혼합물은 다음 펠렛 및 솔루션에 남아있는 작은, 산성 수용성 조각을 형성 소화되지 않은 기판과 centrifuged 있습니다. 뜨는은 해소되고 FITC - 라벨 조각의 형광이 측정됩니다. 설명 키트 절차는 약 0.5 μg / ML (트립신의 5 NG 분석 추가)의 농도에 트립신 프로 테아제 컨트롤을 검색합니다. 이 감도는 최대 24 시간, 긴 배양 시간 증가 수 있습니다. 분석은 microcentrifuge 튜브에서 수행하고 절차 cuvettes 또는 multiwell 판을 사용하여 형광 검출을 위해 제공됩니다.

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

단백 분해 효소 형광 탐지 키트는 fluorometry를 사용하는 프로 테아제 활동의 측정을 위해 설계되었습니다. 그것은 또한 프로 테아제 오염 성분의 검출에 적합합니다. 방법은 FITC - 카제인 표시 기판의 독자적인 배합의 proteolytic hydroysis을 기반으로합니다.

단백 분해 효소의 활동을 확인하는 프로 테아제 형광 검출 키트의 사용

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 조회수.  Sigma Aldrich. 단백 분해 효소 형광 탐지 키트는 fluorometry를 사용하는 프로 테아제 활동의 측정을 위해 설계되었습니다. 그것은 또한 프로 테아제 오염 성분의 검출에 적합합니다. 방법은 FITC - 카제인 표시 기판의 독자적인 배합의 proteolytic hydroysis을 기반으로합니다.

비디오: 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.

Qiagen Miniprep 키트를 사용하여 세균의 플라스미드 식민지에서 DNA를 정화

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

연구

생물학

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.

광학 이미징과 비침습 감지를위한 형광 염료와 함께 줄기 세포를 라벨링

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.

시그마의 비 특정 프로 테아제의 활동 분석 - 카제인 기판으로

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.

효소 아세테이트 키나제의 아세테이트 - 형성 활동의 직접적인 탐지

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.

도입 단백 분해 효소와 산 촉매 라벨링 워크 플로<sup&gt; 18</sup&gt; O 강화 물

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.

Langerhans 절연의 마우스 작은 섬은 정화의 Collagenase와 중성 단백 분해 효소의 조합을 사용하여

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.

형광 단백질 분해 효소 보호를 사용하여 막 단백질 토폴로지를 결정 (FP​​P)

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.

사용<em&gt; 전의 VIVO</em&gt; 전체 기관 이미징 및 정량 조직 조직학는 형광 표지 된 분자의 생물 분포를 확인하는 방법

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.

젖 산 효소 격리 분석 결과-간단 하 고 신뢰할 수 있는 결정 하는 방법을 포유류 세포에서 대량 Autophagic 격리 활동

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.

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