이 작품은 프로젝트 Z2 - 9229 및 프로그램 P2 - 0249와 슬로베니아어 연구 기관에 의해 지원되었다. 이 비디오는 &quot;Electroporation 기반 기술 및 트리 트먼트&quot;류블랴나 대학, 슬로베니아에서 전기 공학부에 의해 biannually 조직 과학 워크샵 및 대학원 과정에 대한 보충 자료를 나타냅니다.

부 I : 예비 단계 <ol><li> 본 실험 중국 햄스터 난소 세포 라인 (CHO - K1)이 사용됩니다. 세포 8% 소태아 혈청 보충 햄 - F12 문화 매체 ~ 0.7x10 5 셀 / ML, 0.15 MG / ML에 실험실 - 테크 II 챔버 (2 우물, 4cm 2 각) (Nalge Nunc, 독일)에 비쳐지게 만들려합니다 L - 글루타민, 16 MG / ML gentamicin (시그마 - 알드리치, Steinheim, 독일에서), 200 단위 / ML Crystacillin (Pliva, 자그레브, 크로아티아), 그리고 37 5 % CO 2에서 incubated ° C. 또는 세포는 또한 polylysine 같은 휴대폰 접착제와 코팅 # 1 유리 커버 풀렸는데 (두께는 0.13-0.16 ㎜)에 도금 수 있습니다. </li><li> 그들의 문화 매체에있는 세포를 품어. 여전히 약 구형 있지만, 단호하게 자신의 막의 작은 부분으로 표면에 부착된 2-4시간의 수율 세포를 지속 인큐베이션. 또는 부화 16~20시간 후 세포가 완전히 표면에 붙어 더 복잡한 모양을 가지고 있지만, 대부분은 여전히​​ 분리되지 않습니다됩니다. </li><li> 원본 Invitrogen의 약병에 든 색소 5 밀리그램 DMSO의 843 μl (시그마 - 알드리치, Steinheim, 독일)를 추가하여 DI - 8 - ANEPPS (Invitrogen, 유진, 오레곤, 미국)의 10 MM 주식 솔루션을 준비합니다. 주식 솔루션은 몇 개월 4 ° C에서 냉장고에 저장할 수 있습니다. DMSO의 크리스탈 녹이신 때까지 실험을 시작하기 전에, 솔루션을 따뜻하게. </li><li> 일부 세포 라인은 세포막에 염료 설립을 쉽게하기 pluronic의 사용이 필요할 수 있습니다. Pluronic는 20 % DMSO (F - 127, Invitrogen, 유진, 오레곤, 미국)의 주식 솔루션 또는 DMSO에 pluronic을 용해하여 준비할 수 같은 농도의 주식 솔루션에 구입하실 수 있습니다. pluronic 재고 솔루션은 상온에서 저장할 수 있습니다. </li></ol> 부 II : DI - 8 - ANEPPS으로 세포 중 <ol><li> 10 MM DI - 8 - ANEPPS 및 최소 필수 매체 SMEM의 스피너 수정 1 ML (칼슘 고갈 버전) (중간 M8167이나 M4767, 시그마 - 알드리치, Steinheim, 독일 20 % pluronic 2.5 μl 3 μl를 혼합 1.5 ML Eppendorf 튜브에). 이것은 약 30 μm의 DI - 8 - ANEPPS 및 pluronic의 0.05 %를 포함하는 &quot;로딩 솔루션&quot;을 산출. 다른 세포 유형에 대한 그들의 고유 문화 미디어는 대신 SMEM에 사용할 수 있습니다. </li><li> 로딩 솔루션 연구소 - 테크 챔버에있는 문화 매체를 교체합니다. 4 10 분 냉장고에 챔버를 전송 ° C. 이 온도에서 플라즈마 막 통해 염료의 국제화가 크게 저해합니다. </li><li> 얼룩 후, 부드럽게 순수 SMEM와 함께 여분의 염료를 두 세번을 씻는다. </li><li> 챔버에 SMEM 1.5 ML를 남겨주세요. </li></ol> 파트 III : 실험 및 이미지 수집 <ol><li> 세포는 기름 침지 목표 (x63, NA 1.4), 단색 (폴리 크롬 IV, Visitron, 독일) 및 냉각 CCD 카메라 (VisiCam 장착된 형광 현미경 (우리의 경우에는 자이스 혈구 AxioVert 200 자이스 혈구, 독일)를 사용하여 관찰 1280, Visitron, 독일). 이미지 MetaFluor 7.1.1로 인수하고 (두 분자 디바이스, 다우닝, PA, 미국) MetaMorph 7.1.1로 처리하지만, 다른 유사한 수집 소프트웨어도 사용할 수 있습니다. 490 nm의에 인수 소프트웨어의 여기 파장을 설정하고 예를 들어 ANEPPS에 적합한 배출 필터, 605의 nm의 중심 대역 통과 필터를 (605/55m, 이색성 565 DCXR, 채도, 로킹엄, VT, 미국)를 선택합니다. 단색를 사용할 수없는 경우, 490 nm의 중심 여기 필터를 대신 사용할 수 있습니다. </li><li> 현미경 단계에 세포와 챔버를 놓고, 챔버의 하단에있는 전극을 위치와 펄스 발생기에 연결합니다. 매체는 전극을 커버한다. </li><li> 세포 생존을 유지하고 가열을 줄이기 위해 뛰고있는 충분히 낮은 진폭과 짧은 기간합니다. 본 실험에서는 35 V 진폭 50 MS 기간 (electroporation를 피할 정도로 낮은)과 사각형 펄스는 DC 전압 공급 장치 및 사용자 정의 - 만든 마이크로 프로세서 제어 교환기 장치를 사용하여 생성됩니다. 또는 이러한 33210A (애질런트, 산타 클라라, CA, 미국)와 같은 상업 펄스 발생기는 증폭기 또는 S88 (그라스, 웨스트 워릭, RI, 미국)와 같은 상업 자극기에 연결, 사용할 수 있습니다. 펄스는 전극과 유도 ΔΦ 사이에 약 88 V / cm의 전기장을 만들고, 0.8 mm의 직경과 그들 사이 4mm 거리 두 개의 병렬 PT / IR 와이어 전극에 전달됩니다. 본 실험에 사용되는 전극 사이의 정확한 필드 분포가 다른 [1] 설명되어 있습니다. </li><li> 관심의 세포를 찾습니다. 하나의 전기 펄스 또는 펄스의 시퀀스를 적용합니다. 즉시 펄스 (관리 이미지) 전에, 그리고 펄스 (펄스 이미지) 동안 하나의 각 펄스에 대한 두 개의 형광 이미지를 획득. DI - 8 - ANEPPS의 낮은 반응으로 인해, 형광의 변화는 육안으로 분별하고 명백한 onl 될 어렵습니다처리 후 Y. 펄스 전송은 수집 소프트웨어의 기능 이미지 수집과 동기화해야합니다. 난방 photobleaching 가능한 피하려면, 조명은 펄스의 기간을 제한할 수 있습니다. </li></ol> 파트 IV : 이미지 처리 및 분석 <ol><li> MetaMorph 소프트웨어에서 이미지를 엽니다. 각 펄스 들어, 컨트롤 및 펄스 이미지 모두에서 배경을 뺍니다. </li><li> 셀을 선택하고 그것이 막에 해당되도록 관심 영역을 설정합니다. 제어 및 펄스 이미지를이 지역에 따라 형광 강도를 측정하고 스프레드 시트에 값을 전송합니다. </li><li> 각 펄스의 경우 펄스 데이터 컨트롤 데이터를 빼야하고, 상대 형광 변화를 얻기 위해 제어 데이터에 의해 결과를 나눕니다. 펄스의 시퀀스가​​ 적용되면, 각 펄스에 대한 결정 상대적으로 형광 변화의 값이보다 신뢰할 수있는 측정을 얻으려면 평균 수 있습니다. </li><li> 교정 곡선을 사용하여 ΔΦ에 상대적인 형광 변화를 변환. 이 곡선의 대략 견적은 문학에서 얻은지만, 높은 정확성, 그것은 각각의 특정 설치에 대해 측정되어야 수 있습니다. 세포가이 실험에서 사용되는 설정을위한 형광의 6 % 감소 ΔΦ [2]에 100 MV 증가에 해당합니다. </li><li> 마지막으로, 이러한 엑셀 (마이크로 소프트, 레드 몬드, WA, 미국), 시그마 플롯 (Systat 소프트웨어 주식 회사, 산호세, CA, 미국)와 같은 그래프 소프트웨어 패키지에 상대 아크 길이의 함수로 전압을 계획하거나, 원산지 (OriginLab 공단, 노샘프턴, MA, 미국). 곡선은 또한 이동 평균 필터로 적합한 필터를 사용하여 smoothed 수 있습니다. </li></ol>

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유도 멤브레인 (transmembrane) 전압, ΔΦ의 측정은 전압 - 게이 티드 멤브레인 채널, 액션 잠재적인 전파, 심장 세포 자극, 또는 세포막의 electroporation [3, 4, 5, 6, 연구 등 다양한 실험 설정에 중요한 수 , 7]. 간단한 세포 형태로 ΔΦ analytically 계산하실 수 있습니다. 예를 들어, 구형 세포에 대한, ΔΦ은 전압이 필드 강도와 셀 크기에 비례되는 상태와 멤브레인 따라 코사인 함수를 다음과 슈완의 방정식에 의해 주어집니다 [8, 9]. 더 복잡한 세포 모양의 경우, ΔΦ 코사인에서 상당히 이탈하고 potentiometric 염료 [12, 13, 14, 15]을 사용하여, [2, 10, 11], 또는 실험적으로 컴퓨터를 사용하거나 숫자 결정되어야합니다. 널리 이러한 목적에 사용되는 potentiometric 염료 중 하나는 DI - 8 - ANEPPS (DI - 8 - 부틸 - 아미노 naphthyl - - pyridinium - 프로필 - 산염 에틸렌), 여기 및 발광의 막 전압에 의존 스펙트럼과 빠른 염료입니다 이것은 세포막과 그 가치를 측정하는 ΔΦ의 변형의 비침 투 관찰 수 있습니다. 이 비디오에서는, 우리는 디 - 8 - ANEPPS을 사용하여 ΔΦ 결정을위한 실험 방법을 보여줍니다. 염료는 코네티컷 대학의 교수 레슬리 로우 및 동료 [13, 14]에 의해 개발하고 빠르게 반응 염료의 클래스에 속하는했다. DI - 8 - ANEPPS은 물에 nonfluorescent이며 이것은 세포막의 지질 이중층에 통합하면 강력하게 형광됩니다. 분자내 전하 분포와 염료의 형광 스펙트럼의 프로필과 강도에 해당 변경 사항 변경 ΔΦ 결과에서 변경할 수 있습니다. DI - 8 - ANEPPS의 형광 강도는 ΔΦ의 변화에​​ 비례 다양하며 염료의 반응은 -280 뮤직 비디오에서 250에 이르기까지 다양한 뮤직 비디오 [4, 16] 전압에 대한 선형이다. 이러한 노력도 [17]을보고 있었지만 염료, 고르지 멤브레인 얼룩의 형광 상대적으로 작은 변화, 그리고 염료 국제화는 멤브레인 전압, 예를 들어 자사의 휴식 구성 요소의 절대 측정을위한 DI - 8 - ANEPPS 덜 적합합니다. 그것은, 그러나, 고르기 세포 [4, 5]에 외부 전기 분야에 노출 nonexcitable 세포에서 유도된 막 전압의 발병 [12, 13], 또는 행동 잠재력으로 막 전압에 큰 변화를 측정에 적합합니다. 여기에 적용되지 않지만, DI - 8 - ANEPPS도 있습니다 형광 여기의 ratiometric 측정하여 ΔΦ 결정 [18] 또는 방출 반응의 감도를 증가하고 상기 효과를 감소 [19]. DI - 8 - ANEPPS 얼룩에 막을로서, 그것은 또한 멤브레인 마커로 노골적으로 사용할 수있는 [2]. 염색의 단점 중 하나는 photobleaching하는 경향이있다이다, 그래서 강한 빛에 장시간 노출을 피해야한다고. 염료의 보정이 외부 매체 중 (I) 칼륨 ionophore의 valinomycin과 다른 칼륨 농도의 집합으로 수행 [2,18], 또는 (ii) 패치 - 클램프 전압 클램프 모드에서 [17]. 마지막으로, 구형 세포와 더 복잡한 형태의 세포에 ΔΦ의 측정, 비디오 ΔΦ의 진폭 및 공간적 분포에 세포 형태의 영향을 보여줍니다. 따라서 구형 세포에 대한 ΔΦ 더 복잡한 세포 형태에 대한 ΔΦ의 공간적 분포가 더 복잡한 반면, 슈완의 방정식과 합의에 코사인 인접해 있습니다 [20] ...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>di-8-ANEPPS</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>D-3167</td>
<td>potentiometric fluorescent dye</td>
</tr>
<tr>
<td>pluronic</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>P3000MP</td>
<td>potassium ionophore</td>
</tr>
<tr>
<td>DMSO</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>D2650</td>
<td>&nbsp;</td>
<tr>
<td>SMEM</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>M8167 or M4767</td>
<td>Spinner modification of the Minimum Essential Medium</td>
</tr>
<tr>
<td>Ham-F12</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>N4888</td>
<td>culture medium</td>
</tr>
<tr>
<td>fetal calf serum</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>F4135</td>
<td>&nbsp;</td>
<tr>
<td>L-glutamine</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>G7513</td>
<td>&nbsp;</td>
<tr>
<td>crystacillin</td>
<td>&nbsp;</td>
<td>Pliva</td>
<td>625110</td>
<td>antibiotic</td>
</tr>
<tr>
<td>gentamicin</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>G1397</td>
<td>antibiotic</td>
</tr>
<tr>
<td>Lab-Tek II</td>
<td>&nbsp;</td>
<td>Nalge Nunc</td>
<td>155379</td>
<td>chamber</td>
</tr>
<tr>
<td>DC voltage supply</td>
<td>&nbsp;</td>
<td>Elektro-Automatik</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>microprocessor-controlled switcher</td>
<td>&nbsp;</td>
<td>Custom made</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>electrodes</td>
<td>&nbsp;</td>
<td>Custom made</td>
<td>&nbsp;</td>
<td>Pt/Ir</td>
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measuring the induced membrane voltage with di-8-anepps

외부 전기장에 세포의 위치는 세포막에 걸쳐 전압 그 결과, 지역 요금 재배포 내부와 세포막의 주변에있는 세포의 외부를 일으 킵니다. 이 전압은 유도 멤브레인 전압을 칭했다 (또한 유도 transmembrane 전압 또는 유도 transmembrane 잠재력의 차이)와 ΔΦ으로 표시, 오직 외부 필드가 존재하는만큼 존재한다. 휴식 전압 막에 존재하면, 유도 전압이에 (추가) superimposes. 이러한 DI - 8 - ANEPPS로 potentiometric 형광 염료, 중 하나를 사용하여, 그것은 세포막에 ΔΦ의 변화를 관찰하고 noninvasively의 가치를 측정하는 것이 가능합니다. ΔΦ의 변화에​​ 비례 형광 강도의 변화와 세포막의 지질 이중층에 바인딩된 경우 DI - 8 - ANEPPS 강력히 형광됩니다. 이 비디오는 측정을위한 프로토콜 ΔΦ DI - 8 - ANEPPS를 사용을 보여줍니다 또한 진폭과 ΔΦ의 공간적 분포에서 셀 모양의 영향을 보여줍니다.

Watch this Scientific Journal Video about Measuring the Induced Membrane Voltage with Di-8-ANEPPS at JoVE.com

Measuring the Induced Membrane Voltage with Di-8-ANEPPS

외부 전기장은 세포의 막에 전압을 유도, 유도 멤브레인 전압 (ΔΦ) 칭했다. potentiometric 염색 DI - 8 - ANEPPS을 사용하여, 그것은 noninvasively ΔΦ를 측정할 수 있습니다. 이 비디오는 측정을위한 프로토콜을 보여줍니다 ΔΦ DI - 8 - ANEPPS를 사용합니다.

디 - 8 - ANEPPS으로 유도된 막 전압 측정

Placement of a cell into an external electric field causes a local charge redistribution inside and outside of the cell in the vicinity of the cell ...

measuring-the-induced-membrane-voltage-with-di-8-anepps

Research

JoVE Journal

Biology

17.3K 조회수.  University of Ljubljana. 외부 전기장은 세포의 막에 전압을 유도, 유도 멤브레인 전압 (ΔΦ) 칭했다. potentiometric 염색 DI - 8 - ANEPPS을 사용하여, 그것은 noninvasively ΔΦ를 측정할 수 있습니다. 이 비디오는 측정을위한 프로토콜을 보여줍니다 ΔΦ DI - 8 - ANEPPS를 사용합니다.

비디오: Measuring the Induced Membrane Voltage with Di-8-ANEPPS

Measuring Near Plasma Membrane and Global Intracellular Calcium Dynamics in Astrocytes

The brain contains glial cells.  Astrocytes, a type of glial cell, have long been known to provide a passive supportive role to neurons. However, increasing evidence suggests that astrocytes may also actively participate in brain function through functional interactions with neurons.  However, many fundamental aspects of astrocyte biology remain controversial, unclear and/or experimentally unexplored. One important issue is the dynamics of intracellular calcium transients in astrocytes. This is relevant because calcium is well established as an important second messenger and because it has been proposed that astrocyte calcium elevations can trigger the release of transmitters from astrocytes. However, there has not been any detailed or satisfying description of near plasma membrane calcium signaling in astrocytes. Total internal reflection fluorescence (TIRF) microscopy is a powerful tool to analyze physiologically relevant signaling events within about 100 nm of the plasma membrane of live cells. Here, we use TIRF microscopy and describe how to monitor near plasma membrane and global intracellular calcium dynamics almost simultaneously. The further refinement and systematic application of this approach has the potential to inform about the precise details of astrocyte calcium signaling. A detailed understanding of astrocyte calcium dynamics may provide a basis to understand if, how, when and why astrocytes and neurons undergo calcium-dependent functional interactions.

We describe how to measure near membrane and global intracellular calcium dynamics in cultured astrocytes using total internal reflection and epifluorescence microscopy.

Astrocytes의 플라즈마 막 글로벌 세포내 칼슘 역학 니어 측정

The brain contains glial cells.  Astrocytes, a type of glial cell, have long been known to provide a passive supportive role to neurons. However, ...

연구

생물학

Measuring Plasma Membrane Protein Endocytic Rates by Reversible Biotinylation

Plasma membrane proteins are a large, diverse group of proteins comprised of receptors, ion channels, transporters and pumps.  Activity of these proteins is responsible for a variety of key cellular events, including nutrient delivery, cellular excitability, and chemical signaling.  Many plasma membrane proteins are dynamically regulated by endocytic trafficking, which modulates protein function by altering protein surface expression.  The mechanisms that facilitate protein endocytosis are complex and are not fully understood for many membrane proteins.  In order to fully understand the mechanisms that control the endocytic trafficking of a given protein, it is critical that the protein s endocytic rate be precisely measured.  For many receptors, direct endocytic rate measurements are frequently achieved utilizing labeled receptor ligands.  However, for many classes of membrane proteins, such as transporters, pumps and ion channels, there is no convenient ligand that can be used to measure the endocytic rate.  In the present report, we describe a reversible biotinylation method that we employ to measure the dopamine transporter (DAT) endocytic rate.  This method provides a straightforward approach to measuring internalization rates, and can be easily employed for trafficking studies of most membrane proteins.

Regulated endocytosis governs the cell surface expression levels of the majority of membrane proteins. Here we utilize reducible, membrane impermeant biotinylation reagents to measure the endocytic rate of the dopamine transporter (DAT), a polytopic membrane protein. The method facilitates a straightforward approach to measuring the endocytic rate of most plasma membrane proteins.

가역 Biotinylation에 의해 플라즈마 막 단백질 Endocytic 요금 측정

Plasma membrane proteins are a large, diverse group of proteins comprised of receptors, ion channels, transporters and pumps.  Activity of these proteins ...

Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the lipidic cubic phase or in meso method. The method has been shown to be quite versatile in that it has been used to solve X-ray crystallographic structures of prokaryotic and eukaryotic proteins, proteins that are monomeric, homo- and hetero-multimeric, chromophore-containing and chromophore-free, and alpha-helical and beta-barrel proteins. Recent successes using in meso crystallization are the human engineered beta2-adrenergic and adenosine A2a G protein-coupled receptors. Protocols are presented for reconstituting the membrane protein into the monoolein-based mesophase, and for setting up crystallizations in the manual mode. Additional steps in the overall process, such as crystal harvesting, are to be addressed in future video articles. The time required to prepare the protein-loaded mesophase and to set up a crystallization plate manually is about one hour.

Herein is described the procedure implemented in the Caffrey Membrane Structural and Functional Biology Group to set up manually crystallization trials of membrane proteins in lipidic mesophases.

Lipidic Mesophases를 사용하여 구조 결정을위한 막 단백질 Crystallizing

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the ...

Crystallization of Membrane Proteins in Lipidic Mesophases

Membrane proteins perform critical functions in living cells related to signal transduction, transport and energy transformations, and, as such, are implicated in a multitude of malfunctions and diseases. However, a structural and functional understanding of membrane proteins is strongly lagging behind that of their soluble partners, mainly, due to difficulties associated with their solubilization and generation of diffraction quality crystals. Crystallization in lipidic mesophases (also known as in meso or LCP crystallization) is a promising technique which was successfully applied to obtain high resolution structures of microbial rhodopsins, photosynthetic proteins, outer membrane beta barrels and G protein-coupled receptors. In meso crystallization takes advantage of a native-like membrane environment and typically produces crystals with lower solvent content and better ordering as compared to traditional crystallization from detergent solutions. The method is not difficult, but requires an understanding of lipid phase behavior and practice in handling viscous mesophase materials. Here we demonstrate a simple and efficient way of making LCP and reconstituting a membrane protein in the lipid bilayer of LCP using a syringe mixer, followed by dispensing nanoliter portions of LCP into an assay or crystallization plate, conducting pre-crystallization assays and harvesting crystals from the LCP matrix. These protocols provide a basic guide for approaching in meso crystallization trials; however, as with any crystallization experiment, extensive screening and optimization are required, and a successful outcome is not necessarily guaranteed.

The protocols describe the essential steps for obtaining diffraction quality crystals of a membrane protein starting from reconstitution of the protein in a lipidic cubic phase (LCP), finding initial conditions with LCP-FRAP pre-crystallization assays, setting up LCP crystallization trials and harvesting crystals.

Lipidic Mesophases의 막 단백질의 결정화

Membrane proteins perform critical functions in living cells related to signal transduction, transport and energy transformations, and, as such, are ...

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

막 단백질의 Conformational 역학 검사 현장에서 사이트 이동 형광 라벨링과

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

The Xenopus Oocyte Cut-open Vaseline Gap Voltage-clamp Technique With Fluorometry

The cut-open oocyte Vaseline gap (COVG) voltage clamp technique allows for analysis of electrophysiological and kinetic properties of heterologous ion channels in oocytes. Recordings from the cut-open setup are particularly useful for resolving low magnitude gating currents, rapid ionic current activation, and deactivation. The main benefits over the two-electrode voltage clamp (TEVC) technique include increased clamp speed, improved signal-to-noise ratio, and the ability to modulate the intracellular and extracellular milieu.
Here, we employ the human cardiac sodium channel (hNaV1.5), expressed in Xenopus oocytes, to demonstrate the cut-open setup and protocol as well as modifications that are required to add voltage clamp fluorometry capability.
The properties of fast activating ion channels, such as hNaV1.5, cannot be fully resolved near room temperature using TEVC, in which the entirety of the oocyte membrane is clamped, making voltage control difficult. However, in the cut-open technique, isolation of only a small portion of the cell membrane allows for the rapid clamping required to accurately record fast kinetics while preventing channel run-down associated with patch clamp techniques.
In conjunction with the COVG technique, ion channel kinetics and electrophysiological properties can be further assayed by using voltage clamp fluorometry, where protein motion is tracked via cysteine conjugation of extracellularly applied fluorophores, insertion of genetically encoded fluorescent proteins, or the incorporation of unnatural amino acids into the region of interest1. This additional data yields kinetic information about voltage-dependent conformational rearrangements of the protein via changes in the microenvironment surrounding the fluorescent molecule.

The Xenopus Oocyte Cut-open Vaseline Gap Voltage-clamp Technique With Fluorometry

The cut-open Vaseline gap approach is used to obtain low noise recordings of ionic and gating currents from voltage-dependent ion channels expressed in Xenopus oocytes with high resolution of fast channel kinetics. With minor modification, voltage clamp fluorometry can be coupled to the cut-open oocyte protocol.

<em&gt; Xenopus의</em형광 측정으로&gt; 난자 절개 바셀린 갭 전압 클램프 기술

The cut-open oocyte Vaseline gap (COVG) voltage clamp technique allows for analysis of electrophysiological and kinetic properties of heterologous ion ...

Imaging Plasma Membrane Deformations With pTIRFM

To gain novel insights into the dynamics of exocytosis, our group focuses on the changes in lipid bilayer shape that must be precisely regulated during the fusion of vesicle and plasma membranes. These rapid and localized changes are achieved by dynamic interactions between lipids and specialized proteins that control membrane curvature. The absence of such interactions would not only have devastating consequences for vesicle fusion, but a host of other cellular functions that involve control of membrane shape. In recent years, the identity of a number of proteins with membrane-shaping properties has been determined. What remains missing is a roadmap of when, where, and how they act as fusion and content release progress.
Our understanding of the molecular events that enable membrane remodeling has historically been limited by a lack of analytical methods that are sensitive to membrane curvature or have the temporal resolution to track rapid changes. PTIRFM satisfies both of these criteria. We discuss how pTIRFM is implemented to visualize and interpret rapid, submicron changes in the orientation of chromaffin cell membranes during dense core vesicle (DCV) fusion. The chromaffin cells we use are isolated from bovine adrenal glands. The membrane is stained with a lipophilic carbocyanine dye,1,1&#39;-dioctadecyl-3,3,3&#39;,3&#39;-tetramethylindodicarbocyanine, 4-chlorobenzenesulfonate, or&#160;diD. DiD intercalates in the membrane plane with a &#34;fixed&#34; orientation and is therefore sensitive to the polarization of the evanescent field. The diD-stained cell membrane is sequentially excited with orthogonal polarizations of a 561&#160;nm laser (p-pol, s-pol). A 488&#160;nm laser is used to visualize vesicle constituents and time the moment of fusion. Exocytosis is triggered by locally perfusing cells with a depolarizing KCl solution. Analysis is performed offline using custom-written software to understand how diD emission intensity changes relate to fusion pore dilation.

Polarization-based Total Internal Reflection Fluorescence Microscopy (pTIRFM) enables real-time detection of cell membrane dynamics. This article describes the implementation of pTIRFM for the study of membrane remodeling during regulated exocytosis. The technique is generalizable to other processes in cell biology that directly or indirectly involve changes in membrane shape.

pTIRFM와 이미징 플라즈마 막 변형

To gain novel insights into the dynamics of exocytosis, our group focuses on the changes in lipid bilayer shape that must be precisely regulated during ...

Measuring Glutathione-induced Feeding Response in Hydra

Hydra is among the most primitive organisms possessing a nervous system and chemosensation for detecting reduced glutathione (GSH) for capturing the prey. The movement of prey organisms causes mechanosensory discharge of the stinging cells called nematocysts from hydra, which are inserted into the prey. The feeding response in hydra, which includes curling of the tentacles to bring the prey towards the mouth, opening of the mouth and consequent engulfing of the prey, is triggered by GSH present in the fluid released from the injured prey. To be able to identify the molecular mechanism of the feeding response in hydra which is unknown to date, it is necessary to establish an assay to measure the feeding response. Here, we describe a simple method for the quantitation of the feeding response in which the distance between the apical end of the tentacle and mouth of hydra is measured and the ratio of such distance before and after the addition of GSH is determined. The ratio, called the relative tentacle spread, was found to give a measure of the feeding response. This assay was validated using a starvation model in which starved hydra show an enhanced feeding response in comparison with daily fed hydra.

Here we describe a simple assay for the quantification of the feeding response in hydra induced by the reduced form of glutathione. This assay relies on measuring the distance between the apical end of the tentacle and mouth of hydra.

히드라의 글루타티온 유도 먹이 반응을 측정

Hydra is among the most primitive organisms possessing a nervous system and chemosensation for detecting reduced glutathione (GSH) for capturing the prey. ...

Microperfusion Technique to Investigate Regulation of Microvessel Permeability in Rat Mesentery

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular mechanisms regulating vascular permeability in cultured endothelial cell monolayers and the functional exchange properties of whole microvascular beds. A method to cannulate and perfuse venular microvessels of rat mesentery and measure the hydraulic conductivity of the microvessel wall is described. The main equipment needed includes an intravital microscope with a large modified stage that supports micromanipulators to position three different microtools: (1) a beveled glass micropipette to cannulate and perfuse the microvessel; (2) a glass micro-occluder to transiently block perfusion and enable measurement of transvascular water flow movement at a measured hydrostatic pressure, and (3) a blunt glass rod to stabilize the mesenteric tissue at the site of cannulation. The modified Landis micro-occlusion technique uses red cells suspended in the artificial perfusate as markers of transvascular fluid movement, and also enables repeated measurements of these flows as experimental conditions are changed and hydrostatic and colloid osmotic pressure difference across the microvessels are carefully controlled. Measurements of hydraulic conductivity first using a control perfusate, then after re-cannulation of the same microvessel with the test perfusates enable paired comparisons of the microvessel response under these well-controlled conditions. Attempts to extend the method to microvessels in the mesentery of mice with genetic modifications expected to modify vascular permeability were severely limited because of the absence of long straight and unbranched microvessels in the mouse mesentery, but the recent availability of the rats with similar genetic modifications using the CRISPR/Cas9 technology is expected to open new areas of investigation where the methods described herein can be applied.

The modified Landis technique enables paired measurement of the hydraulic conductivity of individual microvessels in the mesentery of normal and genetically modified rats under control and test conditions using microperfusion techniques. It provides a convenient method to evaluate mechanisms that regulate microvessel permeability and transvascular exchange under physiological conditions.

Microperfusion 기술은 쥐 장간막에서 미세 혈관 투과성의 규정을 조사하기 위해

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular ...

Ground State Depletion Super-resolution Imaging in Mammalian Cells

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to dramatic leaps in our understanding of how cells function. The development and availability of super-resolution microscopy has considerably extended the limits of optical resolution from ~250-20 nm. Biologists are no longer limited to describing molecular interactions in terms of colocalization within a diffraction limited area, rather it is now possible to visualize the dynamic interactions of individual molecules. Here, we outline a protocol for the visualization and quantification of cellular proteins by ground-state depletion microscopy&#160;for fixed cell imaging. We provide examples from two different membrane proteins, an element of the endoplasmic reticulum translocon, sec61&#946;, and a plasma membrane-localized voltage-gated L-type Ca2+ channel (CaV1.2). Discussed are the specific microscope parameters, fixation methods, photo-switching buffer formulation, and pitfalls and challenges of image processing.

This article outlines a protocol for the detection of one or more plasma and/or intracellular membrane proteins using Ground State Depletion (GSD) super-resolution microscopy in mammalian cells. Here, we discuss the benefits and considerations of using such approaches for the visualization and quantification of cellular proteins.

포유류 세포에 있는 바닥 상태 고갈 슈퍼 해상도 영상

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to ...