비행기는 주식이 작품은 건강 그랜트 (GM62916)과 국립 과학 재단 (National Science Foundation) 부여 (MCB0817107)에 대한 국립 연구소에 의해 지원되었다의 유지 보수를 위해 양 V. Lephart 감사합니다.

polytene 염색체 스쿼시 준비에 대해 다음 프로토콜은 요한센 외에서 설명한 절차에서 조정됩니다. (2009). 1. 셋째 instar Drosophila의 애벌레의 문화 고품질 스쿼시 준비를위한 최적의 polytene 염색체를 얻기 위해서는, 솟아 culturing 조건 (즉, 주위 20 달걀 누워 매일 새로운 병에 표준 4 &quot;비행 병 및 변경에 여성 파리). 선택 뚱보 개인 필수입니다 그들은 아직도 방황하고 있지만 pupation하기 직전하는 동안 세번째 instar의 유충을 등반의 첫 번째 작물에서. 21 우리는 정기적으로 문화 ° C 있지만 18 ° C 예를 들어 같은 몇 가지 목적을 위해 더 적합 있습니다 오르게 염색체를 얻을 것이다 때 밴드 / interband 지역은 높은 해상도 시각해야합니다. 2. Polytene 스쿼시 자료 <ul><li> 드루먼드 해부의 포셉 (2) </li><li> 페트리 요리 (60 X 15mm) </li><li> 프로스트 현미경 슬라이드 (피셔 번호 12-544-3) (폴리 - 라이신 - 코팅) </li><li> 22 X 22mm 번호 15 커버 전표 (피셔 번호 12 520B) (Sigmacote와 코팅, 시그마 # SL2) </li><li> 22 X 40mm 번호 15 커버 전표 (피셔 번호 12 - 530B) </li><li> 김 - 쳐줍니다 </li><li> 20X 목적으로 위상 대비 현미경 </li><li> 소형 듀어 (예 : 진공 플라스크 또는 보온병 병) </li><li> 롱 포셉 </li><li> 면도날 </li><li> 코플린 잘 </li><li> 고무 메이드 또는 유리 섬유 복합 트레이 (또는 이와 동등한 sealable 트레이) </li><li> Parafilm (22mm 사각형으로 절단) </li><li> 175g의 무게 </li></ul> 3. Fixatives 및 솔루션 <ol><li> 배 포름 알데히드 주식. 0.74 G.의 새로운 솔루션을 준비 DH 2 O의 4.0 ML과 1N 코 28 μl로 paraformaldehyde. 65 따뜻한 ° C paraformaldehyde를 해산하고 얼음에 저장합니다. </li><li> 정착액 1. 배 포름 알데히드 주식의 0.5 ML 10X PBS, 50μl 트리톤 X - 100, 3.45 ML DH 2 O 및 1.0 ML의 새로운 솔루션을 준비합니다. 트리톤 X - 100을 분산하고 1 시간 이내에 사용하는 웜. </li><li> 정착액 2. 1.5 ML DH 2 O, 2.5 ML 빙초산, 1 시간 이내에 배 포름 알데히드 재고 및 사용에서 1.0 ML의 새로운 솔루션을 준비합니다. </li><li> Lactoacetic 산성 솔루션입니다. 1 ML 젖산, 2 ML DH 2 O, 3 ML 초산의 솔루션을 준비합니다. </li></ol> 4. Polytene의 염색체 스쿼시 준비 : <ol><li> 물을 애벌레를 씻어 해부를위한 조직 문화 접시에있는 PBS로 전송할 수 있습니다. </li><li> , 입 포셉 한 쌍의와 후크의 팁을 파악 다른 페어와 방법 아래의 2 / 3에 대한 본문을 보유하고 입으로 당겨 침샘이 노출되도록 후크. 타액 두뇌에서 분비와 눈 - antennal 디스크를 분리하고,이 선들의 지방 신체 및 기타 관련 조직 멀리 해부하다. </li><li> 두 잘 우울증 슬라이드 잘 첫 번째와 정착액 2 200-300 microliters 잘 두 정착액 1 200 microliters를 추가합니다. 물론 하나의 정착액 1 번에 침샘 중 하나 쌍을 전송하고 일반적으로 1-2분에 대한 대상 에피토프에 필요한 시간의 금액에 대해 숙고하다. (참고 : 같은 히스톤 수정의 대부분 같은 epitopes가 5 분 이상 고정이 필요할 수 있습니다.) </li><li> 사용 포셉 잘 2 정착액 2 침샘을 전송하고 2 분 동안 품어. </li><li> 깨끗한 Sigmacoted coverslip에 Lactoacetic 산성 솔루션의 10-30 μl에 분비를 전송합니다. 부드럽게 coverslip에 polylysine - 코팅 현미경 슬라이드를 낮추고,이 선들은 슬라이드와 coverslip 사이 있도록 수직 압력을 적용하지 않고, coverslip을 선택합니다. 즉시 조심스럽게 한쪽 가장자리에 집게로 coverslip 쥐고하여 확산 세포 용해와 염색체를 촉진 그리고 부드럽게 조직에 대한 수직 압력을 최소화하기 위해 노력하고, 약간 앞뒤로 그것을 이동. 또는 부드럽게 앞뒤로 coverslip을 이동에도 지우개 연필의 측면이나 손가락을 사용합니다. 앞뒤로 coverslip 이동에 지연 그들이 Lactoacetic 산성 용액에 노출시보다 엄격한되는 경향이 있으므로 염색체 무기의 확산 감소 가능성입니다. 솔루션의 혼돈 스럽 것은 종종 세포 분리의 좋은 표시이다. 부드럽게 몇 번 염색체 확산을 지원 또한 연필의 지우개 측면 (수직 압력을 피하기 위해) obliquely coverslip을 수 도청. </li><li> 즉시 염색체가 잘 확산한지 여부를 확인할 20 40X 목적으로 위상 대비 현미경으로 조직을 검사합니다. </li><li> 확산 염색체가 충분하면 깨끗한 김 - 잎사귀의 스택에 다운 coverslip 측면으로 슬라이드를 설정합니다. 플레이스 두 번째 위에 김 - 닦아 및 전단 것이다 coverslip의 수평 운동을 피하 coverslip가 위치하고 단단히 눌러 어디 위에 엄지를 배치하여 염색체를 평평하게염색체. </li><li> 준비가 용도에 적합한지 확인하기 위해 현미경으로 다시 슬라이드를 검사합니다. 염색체가 부수에 따라 크게 이동하는 경우 후속 준비에 Lactoacetic 산성 솔루션의 적은 볼륨을 사용합니다. 적합한 슬라이드 충분한 번호가 얻은 때까지 반복 4.1-4.8 단계를 반복합니다. 슬라이드의 coverslip에 175g 무게를 놓습니다. </li><li> 액체 질소로 작은 듀어 (예, 보온병 병) 채웁니다. 긴 집게를 사용하여 액체 N 2로 수영 슬라이드 끓는 멈출 때까지, 슬라이드를 제거하고 즉시 coverslip을 뒤집기 한 모퉁이에 깨끗한 면도날의 가장자리를 사용합니다. 4 PBS와 코플린있는 항아리에 슬라이드가 즉시 사용할 수있다면, 장소 ° C. 그렇지 않으면 95 % 에탄올로 가득 코플린 병에 슬라이드를 수집합니다. 서리가 조직 슬라이드에 준수하여 coverslip로 남아 않았 확인 멀리 sublimed 일단 coverslip을 검사합니다. 모든 슬라이드는 PBS (몇 시간 이내에 사용하기 위해) 또는 에탄올 (장기 저장을 위해) 중 하나에 저장된 때까지 슬라이드의 나머지와 함께 반복합니다. </li></ol> 대표 결과 고품질 polytene 스쿼시 준비를위한 첫 번째 중요한 단계는 큰 침샘과 핵 지방 애벌레를 성장하는 것입니다. 두 번째는 연습이 걸릴 수 있습니다 확산 과정과 좋은 기술입니다. 개선 확산 성공 한 팁은 부수 단계에서 lactoacetic 산성 솔루션의 최소한의 볼륨을 찾을 수 있습니다. 이것은 멀리 염색체 팔이 씻을 수있는 과도한 스트리밍 군대를 생성하지 않고 염색체의 확산 충분한을 촉진합니다. 또한 Lactoacetic 산성 용액에 노출 때 그들이 더 엄격한 해짐에 따라 앞뒤로 coverslip 이동에 지연이 염색체 무기의 확산 줄일 수 있다고 지적 가치가있다. 모든 것들이 잘되면, 수많은 잘 확산 polytene 염색체가 있어야합니다. 그림 1은 두 초록색으로 interband 지역에 대한 마커와 염료로 분류, 이러한 준비의 예를 보여주는 얼룩 파란색 줄무늬 지역. 부족 확산을 얻을 경우, 염색체는 그림 2에 표시된 작은 공처럼 보이게됩니다. 반면에, 너무 많은 전이가 발생하는 경우, chomosomes 너무 얇고 확장이나 그림 3과 같이 작은 조각으로 조각난 경우에 따라 것입니다. <img src="/files/ftp_upload/1748/1748fig1.jpg" alt="그림 1" /> 그림 1. Polytene 스쿼시 준비 더블 JIL - 1 히스톤 H3S10 키나제 (빨간색)와 Hoechst (파란색)에 항체으로 표시. <img src="/files/ftp_upload/1748/1748fig2.jpg" alt="그림 2" /> 부족 확산과 그림 2. Polytene 스쿼시. <img src="/files/ftp_upload/1748/1748fig3.jpg" alt="그림 3" /> 그림 3. 너무 확산과 Polytene 호박.

<ol>
	<li>Cai, W., Bao, X., Deng, H., Jin, Y., Girton, J., Johansen, J., Johansen, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RNA+polymerase+II-mediated+transcription+at+active+loci+does+not+require+histone+H3S10+phosphorylation+in+Drosophila.">RNA polymerase II-mediated transcription at active loci does not require histone H3S10 phosphorylation in Drosophila.</a> Development. 135, 2917-2925 (2008).</li><li>DiMario, P., Rosby, R., Cui, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+visualization+of+GFP-fusion+proteins+on+polytene+chromosomes.">Direct visualization of GFP-fusion proteins on polytene chromosomes.</a> Dros. Inf. Serv. 89, 115-118 (2006).</li><li>Johansen, K. M., Cai, W., Deng, H., Bao, X., Zhang, W., Girton, J., Johansen, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+for+studying+transcription+and+epigenetic+chromatin+modification+in+Drosophila+polytene+chromosome+squash+preparations+using+antibodies.">Methods for studying transcription and epigenetic chromatin modification in Drosophila polytene chromosome squash preparations using antibodies.</a> Methods. 48, 387-397 (2009).</li></ol>

기존 스쿼시 고정 프로토콜의 아세트산과 젖산 지방산의 포함은 확산 interband 해상도와 염색체 팔 모두를 용이하게하지만, 불행히도 일부 epitopes이 치료를 생존하지 않습니다. 이러한 에피토프의 예는 H3S10ph입니다 (까 외., 2008). 산성 치료 또한 GFP - 태그 단백질, DiMario 외의 고유의 형광을 꺼버리는 그 단점을 가지고 있기 때문에. (2006) 최근에 산성에 민감한 epitopes의 항체 검출을위한 ...

preparation of drosophila polytene chromosome squashes for antibody labeling

Drosophila 오래 염색질 구조와 셋째 instar의 애벌레의 거대한 침샘 polytene 염색체에서 제공하는 cytological 장점으로 인해 유전자 규제 사이의 관계를 공부 좋아하는 모델 시스템을하고 있습니다. 이 조직에서 염색체는 약 1000 사본을 상승을주는 세포 분열의 부재에서 복제의 많은 순찰을 받고있다. DNA는 특정 단백질의 지방화로 염색질 형태를 상호 연관시킬 수있는 독특한 기회를 제공합니다 크게 확대 염색체의 결과로 각 replicative 사이클 이후에 정렬 남아있다. 따라서, 다른 유전자에 존재하는 epigenetic 수정 정의 및 전사 프로세스의 여러 단계에서 관심이 높은 수준이되었습니다. 이러한 연구에 중요한 도구가 효소에 항체와 polytene 염색체의 라벨, 전사 인자, 또는 관심 히스톤 수정입니다. 이 비디오 프로토콜은 항체 라벨에 대한 Drosophila polytene 염색체를 준비 요한센 실험실에서 사용되는 스쿼시 기술을 보여줍니다.

Watch this Scientific Journal Video about Preparation of Drosophila Polytene Chromosome Squashes for Antibody Labeling at JoVE.com

Preparation of Drosophila Polytene Chromosome Squashes for Antibody Labeling

이 비디오 프로토콜은 준비하는 요한센 실험실에서 사용되는 스쿼시 기법을 보여줍니다<em&gt; Drosophila</em항체 라벨에 대한&gt; polytene 염색체.

항체 라벨링에 대한 Drosophila Polytene의 염색체 Squashes의 준비

Drosophila has long been a favorite model system for studying the relationship between chromatin structure and gene regulation due to the cytological ...

preparation-drosophila-polytene-chromosome-squashes-for-antibody

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31.0K 조회수.  Iowa State University. 이 비디오 프로토콜은 준비하는 요한센 실험실에서 사용되는 스쿼시 기법을 보여줍니다 Drosophila polytene 염색체.

비디오: Preparation of Drosophila Polytene Chromosome Squashes for Antibody Labeling

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos and their dechorionation using bleach are demonstrated. Using a glass pipet attached to a mouth suction tube, we illustrate the removal of all cells from single embryos. The method for dispersing cells from each embyro into a small (5 l) drop of medium on an uncoated glass coverslip is demonstrated. A view through the microscope at 1 hour after plating illustrates the preferred cell density. Most of the cells that survive when grown in defined medium are neuroblasts that divide one or more times in culture before extending neuritic processes by 12-24 hours. A view through the microscope illustrates the level of neurite outgrowth and branching expected in a healthy culture at 2 days in vitro. The cultures are grown in a simple bicarbonate based defined medium, in a 5% CO2 incubator at 22-24&deg;C. Neuritic processes continue to elaborate over the first week in culture and when they make contact with neurites from neighboring cells they often form functional synaptic connections. Neurons in these cultures express voltage-gated sodium, calcium, and potassium channels and are electrically excitable. This culture system is useful for studying molecular genetic and environmental factors that regulate neuronal differentiation, excitability, and synapse formation/function.

This video demonstrates the preparation of primary neuronal cultures from midgastrula stage Drosophila embryos. Views of live cultures show cells 1 hour after plating and differentiated neurons after 2 days of growth in a bicarbonate-based defined medium. The neurons are electrically excitable and form synaptic connections.

Midgastrula 스테이지 Drosophila의 태아에서 문화의 연결 준비

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos ...

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생물학

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

의 준비 Drosophila</em공중 드롭 프로토콜을 사용하여 라이브 영상을위한> 태아

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Fluorescent Labeling of Drosophila Heart Structures

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells that align along each side of the heart wall, supportive alary muscles and, in adults, a layer of ventral longitudinal muscle cells. The contractile fibers house conserved constituents of the muscle cytoarchitecture including densely packed bundles of myofibrils and cytoskeletal/submembranous protein complexes, which interact with homologous components of the extracellular matrix. Here we describe a protocol for the fixation and the fluorescent labeling of particular myocardial elements from the hearts of dissected larvae and semi-intact adult Drosophila. Specifically, we demonstrate the labeling of sarcomeric F-actin and of &alpha;-actinin in larval hearts. Additionally, we perform labeling of F-actin and &alpha;-actinin in myosin-GFP expressing adult flies and of &alpha;-actinin and pericardin, a type IV extracellular matrix collagen, in wild type adult hearts. Particular attention is given to a mounting strategy for semi-intact adult hearts that minimizes handling and optimizes the opportunity for maintaining the integrity of the cardiac tubes and the associated tissues. These preparations are suitable for imaging via fluorescent and confocal microscopy. Overall, this procedure allows for careful and detailed analysis of the structural characteristics of the heart from a powerful genetically tractable model system.

Fluorescent Labeling of Drosophila Heart Structures

Here we describe a basic protocol for fluorescent labeling of different elements of heart tubes from larva and adult Drosophila melanogaster. These specimens are well-suited for imaging via fluorescent or confocal microscopy. This technique permits detailed structural analysis of the features of the hearts from a powerful model organism.

의 형광 라벨링 Drosophila 심장 구조

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells ...

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

의 전자 현미경을위한 배아의 준비 Drosophila 태아의 심장 관

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

Live Dissection of Drosophila Embryos: Streamlined Methods for Screening Mutant Collections by Antibody Staining

Drosophila embryos between stages 14 and 17 of embryonic development can be readily dissected to generate "fillet" preparations. In these preparations, the central nervous system runs down the middle, and is flanked by the body walls. Many different phenotypes have been examined using such preparations. In most cases, the fillets were generated by dissection of antibody-stained fixed whole-mount embryos. These "fixed dissections" have some disadvantages, however. They are time-consuming to execute, and it is difficult to sort mutant (GFP-negative) embryos from stocks in which mutations are maintained over GFP balancer chromosomes. Since 2002, our group has been conducting deficiency and ectopic expression screens to identify ligands for orphan receptors. In order to do this, we developed streamlined protocols for live embryo dissection and antibody staining of collections containing hundreds of balanced lines. We have concluded that it is considerably more efficient to examine phenotypes in large collections of stocks by live dissection than by fixed dissection. Using the protocol described here, a single trained individual can screen up to 10 lines per day for phenotypes, examining 4-7 mutant embryos from each line under a compound microscope. This allows the identification of mutations conferring subtle, low-penetrance phenotypes, since up to 70 hemisegments per line are scored at high magnification with a 40X water-immersion lens.

Live Dissection of Drosophila Embryos: Streamlined Methods for Screening Mutant Collections by Antibody Staining

We describe a streamlined protocol for generating "fillet" preparations of Drosophila embryos of specific genotypes. This protocol allows efficient execution of a variety of genetic screens. It also allows excellent visualization of structures in the late embryo.

라이브 해부 Drosophila 태아 : 항체 스테 이닝에 의해 돌연변이 컬렉션을 검사 늘씬한 방법

Drosophila embryos between stages 14 and 17 of embryonic development can be readily dissected to generate "fillet" preparations.  In these preparations, ...

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three preparations of the brain and visual system that cover the range from the developing eye disc-brain complex in the developing pupae to individual eye and brain dissection from adult flies. All protocols are optimized for the live culture of the preparations. However, we also present the conditions for fixed tissue immunohistochemistry where applicable. Finally, we show live imaging conditions for these preparations using conventional and resonant 4D confocal live imaging in a perfusion chamber. Together, these protocols provide a basis for live imaging on different time scales ranging from functional intracellular assays on the scale of minutes to developmental or degenerative processes on the scale of many hours.

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

This protocol describes three Drosophila preparations: 1) adult brain dissection, 2) adult retina dissection and 3) developing eye disc- brain complexes dissection. Emphasis is laid on special preparation techniques and conditions for live imaging, although all preparations can be used for fixed tissue immunohistochemistry.

개발 및 성인의 준비 Drosophila 두뇌와 라이브 영상을위한 Retinae

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three ...

Preparation of Drosophila S2 cells for Light Microscopy

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature and genome sequence database. Cultured Drosophila S2 cells, the product of disassociated 20-24 hour old embryos1, possess all these properties. Consequently, S2 cells are extremely well-suited for the analysis of cellular processes, including the discovery of the genes encoding the molecular components of the process or mechanism of interest. The features of S2 cells that are most responsible for their utility are the ease with which they are maintained, their exquisite sensitivity to double-stranded (ds)RNA-mediated interference (RNAi), and their tractability to fluorescence microscopy as either live or fixed cells.
S2 cells can be grown in a variety of media, including a number of inexpensive, commercially-available, fully-defined, serum-free media2. In addition, they grow optimally and quickly at 21-24&deg;C and can be cultured in a variety of containers. Unlike mammalian cells, S2 cells do not require a regulated atmosphere, but instead do well with normal air and can even be maintained in sealed flasks.
Complementing the ease of RNAi in S2 cells is the ability to readily analyze experimentally-induced phenotypes by phase or fluorescence microscopy of fixed or live cells. S2 cells grow in culture as a single monolayer but do not display contact inhibition. Instead, cells tend to grow in colonies in dense cultures. At low density, S2 cultures grown on glass or tissue culture-treated plastic are round and loosely-attached. However, the cytology of S2 cells can be greatly improved by inducing them to flatten extensively by briefly culturing them on a surface coated with the lectin, concanavalin A (ConA)3. S2 cells can also be stably transfected with fluorescently-tagged markers to label structures or organelles of interest in live or fixed cells. Therefore, the usual scenario for the microscopic analysis of cells is this: first, S2 cells (which can possess transgenes to express tagged markers) are treated by RNAi to eliminate a target protein(s). RNAi treatment time can be adjusted to allow for differences in protein turn-over kinetics and to minimize cell trauma/death if the target protein is important for viability. Next, the treated cells are transferred to a dish containing a coverslip pre-coated with conA to induce cells to spread and tightly adhere to the glass. Finally, cells are imaged with the researcher's choice of microscopy modes. S2 cells are particularly good for studies requiring extended visualization of live cells since these cells stay healthy at room temperature and normal atmosphere.

Preparation of Drosophila S2 cells for Light Microscopy

Drosophila Schneider (S2) cells are an increasingly popular system for the discovery and functional analysis of genes. Our goal is to describe some of the microscopic techniques that make S2 cells such an increasingly important experimental system.

의 준비 Drosophila</em라이트 현미경을 위해> S2 세포

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature ...

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a plethora of mutant strains and technical tricks have been developed to allow sophisticated questions to be asked and answered in a reasonable amount of time. Fundamental insight into the interplay of components of all known major signaling pathways has been obtained in forward and reverse genetic Drosophila studies. The fly eye has proven to be exceptionally well suited for mutational analysis, since, under laboratory conditions, flies can survive without functional eyes. Furthermore, the surface of the insect eye is composed of some 800 individual unit eyes (facets or ommatidia) that form a regular, smooth surface when looked at under a dissecting microscope. Thus, it is easy to see whether a mutation might affect eye development or growth by externally looking for the loss of the smooth surface ('rough eye' phenotype; Fig. 1) or overall eye size, respectively (for examples of screens based on external eye morphology see e.g.1). Subsequent detailed analyses of eye phenotypes require fixation, plastic embedding and thin-sectioning of adult eyes.
The Drosophila eye develops from the so-called eye imaginal disc, a bag of epithelial cells that proliferate and differentiate during larval and pupal stages (for review see e.g. 2). Each ommatidium consists of 20 cells, including eight photoreceptors (PR or R-cells; Fig. 2), four lens-secreting cone cells, pigment cells ('hexagon' around R-cell cluster) and a bristle. The photoreceptors of each ommatidium, most easily identified by their light sensitive organelles, the rhabdomeres, are organized in a trapezoid made up of the six "outer" (R1-6) and two "inner" photoreceptors (R7/8; R8 [Fig. 2] is underneath R7 and thus only seen in sections from deeper areas of the eye). The trapezoid of each facet is precisely aligned with those of its neighbors and the overall anteroposterior and dorsoventral axes of the eye (Fig. 3A). In particular, the ommatidia of the dorsal and ventral (black and red arrows, respectively) halves of the eye are mirror images of each other and correspond to two chiral forms established during planar cell polarity signaling (for review see e.g. 3).
The method to generate semi-thin eye sections (such as those presented in Fig. 3) described here is slightly modified from the one originally described by Tomlinson and Ready4. It allows the morphological analysis of all cells except for the transparent cone cells. In addition, the pigment of R-cells (blue arrowheads in Fig. 2 and 3) can be used as a cell-autonomous marker for the genotype of a R-cell, thus genetic requirements of genes in a subset of R-cells can readily be determined5,6.

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

A standard approach to prepare adult Drosophila eyes for semi-thin sectioning and light microscopic analysis is presented here. The protocol can be used for gross morphological analysis of eye defects, or with the indicated adjustments can be used to determine genetic requirements of genes in specific cell types of the eye (e.g. clonal analysis of photoreceptors) or for electron microscopic analysis.

성인의 준비 Drosophila</em씬 Sectioning 및 현미경 분석을위한> 눈

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a ...

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

생활에서 보여주는 동적 Kinetochore 단백질 행동과 염색체 모션으로 Mitotic 검사 점 유학<em&gt; Drosophila</em&gt; Syncytial 태아

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle ...

Chromosome Preparation From Cultured Cells

Chromosome (cytogenetic) analysis is widely used for the detection of chromosome instability. When followed by G-banding and molecular techniques such as fluorescence in situ hybridization (FISH), this assay has the powerful ability to analyze individual cells for aberrations that involve gains or losses of portions of the genome&#160;and rearrangements involving one or more chromosomes. In humans, chromosome abnormalities occur in approximately 1 per 160 live births1,2, 60-80% of all miscarriages3,4, 10% of stillbirths2,5, 13% of individuals with congenital heart disease6, 3-6% of infertility cases2, and in many patients with developmental delay&#160;and birth defects7. Cytogenetic analysis of malignancy is routinely used by researchers and clinicians, as observations of clonal chromosomal abnormalities have been shown to have both diagnostic and prognostic significance8,9.&#160; Chromosome isolation is invaluable for gene therapy and stem cell research of organisms including nonhuman primates and rodents10-13.
Chromosomes can be isolated from cells of live tissues, including blood lymphocytes, skin fibroblasts, amniocytes, placenta, bone marrow, and tumor specimens. Chromosomes are analyzed at the metaphase stage of mitosis, when they are most condensed and therefore more clearly visible. The first step of the chromosome isolation technique involves the disruption of the spindle fibers by incubation with Colcemid, to prevent the cells from proceeding to the subsequent anaphase stage. The cells are then treated with a hypotonic solution and preserved in their swollen state with Carnoy&#39;s fixative. The cells are then dropped on to slides and can then be utilized for a variety of procedures. G-banding involves trypsin treatment followed by staining with Giemsa to create characteristic light and dark bands. The same procedure to isolate chromosomes can be used for the preparation of cells for procedures such as fluorescence in situ hybridization (FISH), comparative genomic hybridization (CGH), and spectral karyotyping (SKY)14,15.

Chromosomes can be isolated from live cells such as lymphocytes or skin fibroblasts, and from organisms including humans or mice. These chromosome preparations can be further utilized for routine G-banding and molecular cytogenetic procedures such as fluorescence in situ hybridization (FISH), comparative genomic hybridization (CGH), and spectral karyotyping (SKY).

배양 세포에서 염색체 준비

Chromosome (cytogenetic) analysis is widely used for the detection of chromosome instability. When followed by G-banding and molecular techniques such as ...