The authors thank Ian Clarke and P. Scott Hefty for the pGFP-SW2 and pASK-GFP-mKate2-L2 plasmids, respectively. We thank Paul Miller for technical assistance and Richard Stephens for other resources. This work was funded by NIH grant AI095603 (KH).

1.代荧光宿主细胞系<ol><li>以2×10 6个细胞/孔，于6孔板天1.板293T细胞（或其它逆转录病毒包装线）为〜75％汇合的第二天。如果需要的话，所用板的孔一式两份每个逆转录病毒。 </li><li>第2天，吸细胞和加2ml新鲜生长培养基（DMEM + 10％FBS + 2mM的L-谷氨酰胺）。转染细胞用5-8微克每逆转录病毒载体（含有GFP）和包装载体的（如，pVSVg）使用Lipofectamine 2000或类似试剂，并按照制造商的说明。 </li><li>孵育细胞用DNA 4-6小时在37℃ 的 CO 2培养箱，并且随后替换培养基用1ml生长培养基中。 </li><li>孵育细胞48小时在37℃ 的 CO 2培养箱中培养。 </li><li>第3天板靶细胞（例如，HeLa细胞）上的6孔板上，10 5个细胞的近似密度/孔，为〜50％汇合的明天。通过在倒置显微镜监测GFP荧光验证在293T细胞阳性转染。 </li><li>第4天收集通过0.45μm的醋酸纤维素过滤器的注射器用吸管和过滤上清上清关闭293T细胞含有逆转录病毒。 </li><li>从HeLa细胞中删除，并添加含有16微克/毫升聚凝胺0.5毫升生长培养基。添加〜0.5毫升过滤逆转录病毒的细胞逐滴孵育细胞在37℃ 的 CO 2培养箱24小时。存储任何剩余的反转录病毒（例如，从复制孔）在4℃下。 </li><li>日5.如果复孔被做产生额外的逆转录病毒，重复步骤1.7至连续用剩下的逆转录病毒颗粒感染。 </li><li>天6.通道转染的HeLa细胞中1：1到10厘米培养皿含有选择抗生素（例如，500微克/ ml新霉素的）。 </li><li>等待足够的时间转导的细胞重新生长在选择antibio存在抽动，然后将细胞可通过标准技术来传播，但有选择抗生素的浓度较低（例如，200微克/ ml新霉素的）。 注：细胞也可以被克隆选择亮度理想在这个时候，如果有必要的。 </li></ol> 2代的绿色荧光蛋白表达沙眼衣原体 <ol><li>制备2× 氯化钙缓冲液（20mM的Tris pH值7.4，100mM的氯化钙 2）和4SP缓冲液（0.4M的蔗糖，16毫的 Na 2 HPO 4）。 </li><li>每日1次解冻冷冻C.衣原体股票在冰上，并立即稀释10微升细菌进入50微升2倍氯化钙2缓冲区。加入3微克质粒DNA（例如，帕斯克-GFP-mKate2-L2），拌匀，孵育30分钟，在25℃。 </li><li>在此步骤期间，通过胰蛋白酶McCoy细胞的T 75烧瓶入离心管中制备的宿主细胞。离心细胞悬浮液，在50×g离心5分钟，丢弃上清液。 Resuspen在1× 氯化钙缓冲适当卷D细胞沉淀，得到的4×10 7个细胞/ ml的终浓度。 </li><li>在30分钟温育（来自步骤2.2）后，加入100微升制备McCoy细胞以转化混合物管（总体积200微升）并孵育另外的20分钟，在25℃。加入100微升此转化细胞混合物的单个井从6孔板和覆盖用2ml生长培养基中。 在 CO 2培养箱2天在37℃。 </li><li>天3.裂解McCoy细胞感染C.由沙眼用1毫升的dh 2 O与弯曲1000微升枪头更换培养基和刮细胞。加入1毫升4SP缓冲区，并通过一个27.5摹针通过悬挂〜5次，C的高效细胞裂解，并释放衣原体 。 </li><li>含C的细胞裂解物转移2 ml的衣原体到T-75烧瓶中含有新鲜镀MCC的汇合单层OY细胞;加入8 ml的DMEM（无FBS），并培育2小时，在25℃，以允许℃。衣原体坚持细胞。 </li><li>抽吸细胞，并添加补充有10U / ml青霉素G和1μg/ ml的放线菌酮的新鲜生长培养基中。孵育烧瓶48小时在37℃ 的 CO 2培养箱中培养。 </li><li> 4日通过光学显微镜的细胞被感染和夹杂物含有异常衣原体检查。确定夹杂亮泡在一个标准的光学显微镜，以及异常的机构衣原体夹杂物是大于2微米直径内环形物体。 </li><li>日5.裂解文化通过用2毫升的dh 2 O更换培养基和刮细胞成混悬液。加入2ml 4SP缓冲区，并通过一个27.5摹针通过悬挂〜5倍。 </li><li>用2 ml的细胞裂解物去感染McCoy细胞的新鲜单层在单个孔在6孔板中。孵育细胞裂解物2小时在25℃，吸液和添加新鲜的生长中含有青霉素G和放线菌酮。 </li><li>孵育细胞在37℃ 的 CO 2培养箱中3-5天，这取决于细胞的一般健康状况。 </li><li>重复步骤2.9-2.10一次或多次，根据需要，传代培养进新鲜的孔中一个6孔板直到正常寻找夹杂物（不含异常机构）已经出现。此时使用倒置荧光显微镜来检查C.衣原体表达mKate2（红色）。 </li><li>扩大感染的培养物中产生高滴度衣原体股，例如通过从在T-150瓶中生长感染的McCoy或HeLa细胞收获衣原体转化体。 </li></ol> 3.感染细胞衣原体 <ol><li>板的GFP HeLa细胞上所需的培养容器，用于高分辨率成像为IFU测定和多孔筛例如玻璃底培养皿或室载玻片，或24孔板中。 </li><li>解冻来回含C.禅管衣原体，C。 muridarum或 C. 肺炎在冰上并在HBSS中稀释至感染复数（MOI）的所需多样性，例如MOI = 1。 </li><li>执行基于特定衣原体菌株，将用于静态或离心辅助感染。 <ol><li>对于感染C.衣原体轻型货车血清型L2或C. muridarum，洗的GFP HeLa细胞用HBSS，并在25℃下孵育稀释的细菌2小时。 ，洗板细胞与HBSS，并培育细胞生长培养基37℃的二氧化碳培养箱。 </li><li>对于感染沙眼衣原体血清型D或C.肺炎 ，洗GFP-HeLa细胞与HBSS，并添加稀释细菌细胞。在吊桶式转子连接在台式离心机的多孔板架放置多孔板或玻璃底菜（用胶带多孔板盖子固定）。 </li><li>离心细胞，在900×g离心，在25℃1小时。 </l我&gt; <li>从离心机并置于37℃ 的 CO 2培养箱中取出培养容器1小时。 </li><li>吸细胞的生物安全柜，两次HBSS洗涤细胞，并加入新鲜的生长培养基。位置细胞在37℃ 的 CO 2培养箱中培养。 </li></ol></li></ol> 衣原体夹杂物4.活细胞可视化<ol><li>除去衣原体感染的GFP的HeLa从二氧化碳培养箱细胞用RPMI替换培养基无酚红+ 5％FBS中。 </li><li>摩细胞在倒置荧光显微镜（尼康的Eclipse的Ti-E或类似），使用20X或更高的油价目标。 </li><li>使用图像处理软件（Volocity 6或类似），专注于识别和衣原体感染GFP-HeLa细胞：含有大量的黑洞（ 衣原体包涵体）绿色细胞。 </li><li>马克包含目的使用映像软件衣原体 -感染的细胞区域的XYZ位置（卷ocity 6或类似）。通过手动&#39;加分&#39;，而在XY舞台视图模式下执行此。以这种方式，XYZ坐标值保存为每个标记的位置。 </li><li>在采集设置对话框中，配置软件获得绿色（GFP，HeLa细胞细胞质）和红色（mKate2， 沙眼衣原体 ）通道，并在每个通道的新帧，每5分钟的速度。 </li><li>获取使用协议通过单击“捕捉/记录”按钮上面输入采集的图像序列。 </li></ol> 5.定量的活细胞夹杂物生长参数<ol><li>在期望的时间后感染（如，24，48 HPI）取出衣原体感染GFP-HeLa细胞从孵化器，并安装在一个倒置荧光显微镜。注意：用高NA 20X或40X空气目标是最适合于24孔板，并且一个40X油物镜推荐用于玻片。生长细胞或者基质这个屁股唉。 </li><li>着眼于细胞，其中包裹体是在其最大直径的中平面和屈服清晰的边缘。 </li><li>获取图像用于许多领域中的每个孔中（至少每孔10）;井通常代表复制或实验变量。 </li><li>手动枚举夹杂物的数量，由眼（夹杂物是黑色的隔室中的GFP HeLa细胞），或使用成像软件算法来自动识别和计数的内含物。 <ol><li>发现GFP-HeLa细胞荧光强度（“找对象”命令），并调整基于细胞的相对强度荧光阈强度。 </li><li>使用这些保留仅大于5微米2的大小准则筛选选定的对象。这是“人口1”。 </li><li>应用“填写对象洞”利用“人口1”作为输入的命令。此步骤生成&#39;人口2&#39;。 </li><li>从&#39;P减“人口1”opulation 2“产生积极的标记衣原体夹杂物，被指定为”人口3“。 </li><li>适用尺寸过滤器，以&#39;人口3&#39;（“过滤人口&#39;命令），例如保持对象大于25微米2为在感染的细胞24小时或更长的内含物。对于早期感染倍，如之前18小时，设定尺寸过滤器，以保持物体大于4微米2，因为这些夹杂物要小得多。尺寸过滤结果被指定为“夹杂物”的对象的人口。 </li><li>计算从图像的定量数据，例如，包括数字，夹杂物尺寸，以及夹杂物周，使用图像处理软件。 </li></ol></li></ol>

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作者宣称，他们有没有竞争的财务权益。

在这里，我们描述实验的策略产生荧光宿主细胞进行实时可视化和衣原体夹杂物的分析。这种液泡可视化方法赋予强大能力来照亮，跟踪和定量测定衣原体夹杂物的动态属性跨细胞群或单细胞中随着时间的推移， 衣原体夹杂在荧光蛋白标记的细胞惊人地明确定义，使得它们很容易识别而不需要附加的免疫处理。此外，这种方法是足够敏感的解决签名形态差异，不同的衣原体物...

引起细胞内细菌衣原体种传染病引起全球健康的一大负担，包括性传播疾病，盆腔炎，失明，肺炎和动脉粥样硬化有可能1-4。 衣原体到与宿主细胞相互作用，从一个液泡内的能力（称为包含），是一个重要的决定因素的成功细胞和宿主的感染。列入是一种新型的致病隔室，使衣原体生长和在整个衣原体 5的整个 2-3天的发育循环被动态地修改。衣原体的专性细胞内的性质提出了直接学习列入独特的生物学无数挑战，研究界，尤其如此。一个主要的障碍已经无法有效地可视化或者细胞内沙眼或将其列入荧光方法ES在活细胞。最近发现终于露出来产生绿色荧光蛋白表达C.手段衣原体 6;然而，这一发现还没有导致列入特异性标记。一些技术已经被描述用于细菌和夹杂物7,8的标记，但是它们从缺点，例如非特异性，顷刻和易感性漂白受损。一个重要的发现我们集团成立了一个新的战略，利用绿色荧光蛋白表达的宿主细胞9照明列入。这一战略合理利用纳入膜的固有抗渗性的分子大于520大10。当细胞改造成稳定表达特定的细胞内荧光蛋白（ 如 GFP或mCherry）， 衣原体夹杂物具有显着的清晰度荧光他们完全排除可见。这种倒车影像战略使夹杂物的所有叶绿素即时可视化amydia物种，它可以容易地适用于大多数感兴趣的宿主细胞。由于其效用的演示，这种方法以前用来揭示和定义细胞退出途径衣原体属9。 在这里，我们进一步说明此方法的执行，并且可以被利用来导出关于夹杂物的生长动力学键的定量数据。此外，它可以有效地替代昂贵的基于抗体的枚举的方法和可以串联与其他荧光标记，如mKate2表达衣原体 11被使用。这个强大的工具结合，使生活在宿主细胞内的衣原体包涵体膜的物理性质的探索。

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Lipofectamine 2000</td>
			<td>Invitrogen</td>
			<td>11668</td>
			<td></td>
		</tr>
		<tr>
			<td>Opti-Mem</td>
			<td>Invitrogen</td>
			<td>31985</td>
			<td></td>
		</tr>
		<tr>
			<td>Polybrene</td>
			<td>Sigma</td>
			<td>H9268</td>
			<td></td>
		</tr>
		<tr>
			<td>0.45 &#181;m filters</td>
			<td>Fisher</td>
			<td>09-719D</td>
			<td></td>
		</tr>
		<tr>
			<td>G418</td>
			<td>Invitrogen</td>
			<td>10131</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS</td>
			<td>Invitrogen</td>
			<td>14025</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Invitrogen</td>
			<td>11995</td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI 1640</td>
			<td>Invitrogen</td>
			<td>11875</td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI 1640 w/o phenol red</td>
			<td>Cellgro</td>
			<td>17-105-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin G</td>
			<td>Sigma</td>
			<td>13752</td>
			<td></td>
		</tr>
		<tr>
			<td>Cycloheximide</td>
			<td>Sigma</td>
			<td>C7698</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass bottom dishes</td>
			<td>MatTek</td>
			<td>P35G-1.5-14-C</td>
			<td></td>
		</tr>
		<tr>
			<td>Chamber slides, Lab-Tek II</td>
			<td>Nunc</td>
			<td>154526, 154534</td>
			<td></td>
		</tr>
	</tbody>
</table>

using fluorescent proteins to visualize and quantitate chlamydia vacuole growth dynamics in living cells

The obligate intracellular bacterium Chlamydia elicits a great burden on global public health. C.&#160;trachomatis is the leading bacterial cause of sexually transmitted infection and also the primary cause of preventable blindness in the world. An essential determinant for successful infection of host cells by Chlamydia is the bacterium's ability to manipulate host cell signaling from within a novel, vacuolar compartment called the inclusion. From within the inclusion, Chlamydia acquire nutrients required for their 2-3 day developmental growth, and they additionally secrete a panel of effector proteins onto the cytosolic face of the vacuole membrane and into the host cytosol. Gaps in our understanding of Chlamydia biology, however, present significant challenges for visualizing and analyzing this intracellular compartment. Recently, a reverse-imaging strategy for visualizing the inclusion using GFP expressing host cells was described. This approach rationally exploits the intrinsic impermeability of the inclusion membrane to large molecules such as GFP. In this work, we describe how GFP- or mCherry-expressing host cells are generated for subsequent visualization of chlamydial inclusions. Furthermore, this method is shown to effectively substitute for costly antibody-based enumeration methods, can be used in tandem with other fluorescent labels, such as GFP-expressing Chlamydia, and can be exploited to derive key quantitative data about inclusion membrane growth from a range of Chlamydia species and strains.

表达胞质荧光蛋白（例如，GFP）的哺乳动物细胞可被工程化以使衣原体夹杂照明活，感染的细胞培养物。一旦感染衣原体 ，夹杂物如黑点在宿主细胞中（图1）容易看见。荧光缺乏夹杂物的清晰度可以被利用为夹杂物的整个视和/或处理（ 图1）众多领域的自动识别。一旦已经产生荧光的宿主细胞，一个强大的新工作流启用实验样本中衣原体感染水平的?...

Watch this Scientific Journal Video about Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells at JoVE.com

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

A live cell fluorescent protein based method for illuminating cellular vacuoles (inclusions) containing Chlamydia is described. This strategy enables rapid, automated determination of Chlamydia infectivity in samples and can be used to quantitatively investigate inclusion growth dynamics.

利用荧光蛋白可视化和定量<em&gt;衣原体</em&gt;液泡生长动态的活细胞

The obligate intracellular bacterium Chlamydia elicits a great burden on global public health. C.&#160;trachomatis is the leading bacterial cause of ...

using-fluorescent-proteins-to-visualize-quantitate-chlamydia-vacuole

Research

JoVE Journal

Immunology and Infection

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

7.5K 次观看.  University of Washington. A live cell fluorescent protein based method for illuminating cellular vacuoles (inclusions) containing Chlamydia is described. This strategy enables rapid, automated determination of Chlamydia infectivity in samples and can be used to quantitatively investigate inclusion growth dynamics.

视频: Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

Measuring Calpain Activity in Fixed and Living Cells by Flow Cytometry

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, including signaling, cytoskeletal remodeling, regulation of gene expression, apoptosis and cell cycle progression 1. Calpains have been implicated in many pathologies including muscular dystrophies, cancer, diabetes, Alzheimer's disease and multiple sclerosis 1. Calpain regulation is complex and incompletely understood. mRNA and protein levels correlate poorly with activity, limiting the use of gene or protein expression techniques to measure calpain activity. This video protocol details a flow cytometric assay developed in our laboratory for measuring calpain activity in fixed and living cells. This method uses the fluorescent substrate BOC-LM-CMAC, which is cleaved specifically by calpain, to measure calpain activity. 2 In this video, calpain activity in fixed and living murine 32Dkit leukemia cells, alone or as part of a splenocyte population is measured using an LSRII (BD Bioscience). 32Dkit cells are shown to have elevated activity compared to normal splenocytes.

This article will detail the protocol for measuring calpain activity in fixed and living cells using flow cytometry.

在固定和活细胞，流式细胞仪测量钙蛋白酶活性

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, ...

科研

免疫与感染

Seven Steps to Stellate Cells

Hepatic stellate cells are liver-resident cells of star-like morphology and are located in the space of Disse between liver sinusoidal endothelial cells and hepatocytes1,2. Stellate cells are derived from bone marrow precursors and store up to 80% of the total body vitamin A1, 2. Upon activation, stellate cells differentiate into myofibroblasts to produce extracellular matrix, thus contributing to liver fibrosis3. Based on their ability to contract, myofibroblastic stellate cells can regulate the vascular tone associated with portal hypertension4. Recently, we demonstrated that hepatic stellate cells are potent antigen presenting cells and can activate NKT cells as well as conventional T lymphocytes5. 

 Here we present a method for the efficient preparation of hepatic stellate cells from mouse liver. Due to their perisinusoidal localization, the isolation of hepatic stellate cells is a multi-step process. In order to render stellate cells accessible to isolation from the space of Disse, mouse livers are perfused in situ with the digestive enzymes Pronase E and Collagenase P. Following perfusion, the liver tissue is subjected to additional enzymatic treatment with Pronase E and Collagenase P in vitro. Subsequently, the method takes advantage of the massive amount of vitamin A-storing lipid droplets in hepatic stellate cells. This feature allows the separation of stellate cells from other hepatic cell types by centrifugation on an 8% Nycodenz gradient. The protocol described here yields a highly pure and homogenous population of stellate cells. Purity of preparations can be assessed by staining for the marker molecule glial fibrillary acidic protein (GFAP), prior to analysis by fluorescence microscopy or flow cytometry. Further, light microscopy reveals the unique appearance of star-shaped hepatic stellate cells that harbor high amounts of lipid droplets. 


 Taken together, we present a detailed protocol for the efficient isolation of hepatic stellate cells, including representative images of their morphological appearance and GFAP expression that help to define the stellate cell entity.

Here we describe a method for the isolation of hepatic stellate cells from mouse liver. For stellate cell purification, mouse livers are digested in situ and in vitro by pronase-collagenase treatment prior to density gradient centrifugation. This technique yields highly pure hepatic stellate cells.

星状细胞的七个步骤

Hepatic stellate cells are liver-resident  cells of star-like morphology and are located in the space of Disse between  liver sinusoidal endothelial cells ...

Following Cell-fate in E. coli After Infection by Phage Lambda

The system comprising bacteriophage (phage) lambda and the bacterium E. coli has long served as a paradigm for cell-fate determination1,2. Following the simultaneous infection of the cell by a number of phages, one of two pathways is chosen: lytic (virulent) or lysogenic (dormant)3,4. We recently developed a method for fluorescently labeling individual phages, and were able to examine the post-infection decision in real-time under the microscope, at the level of individual phages and cells5. Here, we describe the full procedure for performing the infection experiments described in our earlier work5. This includes the creation of fluorescent phages, infection of the cells, imaging under the microscope and data analysis. The fluorescent phage is a "hybrid", co-expressing wild- type and YFP-fusion versions of the capsid gpD protein. A crude phage lysate is first obtained by inducing a lysogen of the gpD-EYFP (Enhanced Yellow Fluorescent Protein) phage, harboring a plasmid expressing wild type gpD. A series of purification steps are then performed, followed by DAPI-labeling and imaging under the microscope. This is done in order to verify the uniformity, DNA packaging efficiency, fluorescence signal and structural stability of the phage stock. The initial adsorption of phages to bacteria is performed on ice, then followed by a short incubation at 35&deg;C to trigger viral DNA injection6. The phage/bacteria mixture is then moved to the surface of a thin nutrient agar slab, covered with a coverslip and imaged under an epifluorescence microscope. The post-infection process is followed for 4 hr, at 10 min interval. Multiple stage positions are tracked such that ~100 cell infections can be traced in a single experiment. At each position and time point, images are acquired in the phase-contrast and red and green fluorescent channels. The phase-contrast image is used later for automated cell recognition while the fluorescent channels are used to characterize the infection outcome: production of new fluorescent phages (green) followed by cell lysis, or expression of lysogeny factors (red) followed by resumed cell growth and division. The acquired time-lapse movies are processed using a combination of manual and automated methods. Data analysis results in the identification of infection parameters for each infection event (e.g. number and positions of infecting phages) as well as infection outcome (lysis/lysogeny). Additional parameters can be extracted if desired.

Following Cell-fate in E. coli After Infection by Phage Lambda

This article describes the procedure for preparing a fluorescently-labeled version of bacteriophage lambda, infection of E. coli bacteria, following the infection outcome under the microscope, and analysis of infection results.

在细胞命运<em&gt; E.大肠杆菌</em感染后的噬菌体λ

The system comprising bacteriophage (phage) lambda and the bacterium E. coli has long served as a paradigm for cell-fate determination1,2. Following the ...

Measuring Growth and Gene Expression Dynamics of Tumor-Targeted S. Typhimurium Bacteria

The goal of these experiments is to generate quantitative time-course data on the growth and gene expression dynamics of attenuated S. typhimurium bacterial colonies growing inside tumors. 
We generated model xenograft tumors in mice by subcutaneous injection of a human ovarian cancer cell line, OVCAR-8 (NCI DCTD Tumor Repository, Frederick, MD). 
We transformed attenuated strains of S. typhimurium bacteria (ELH430:SL1344 phoPQ- 1) with a constitutively expressed luciferase (luxCDABE) plasmid for visualization2. These strains specifically colonize tumors while remaining essentially non-virulent to the mouse1. 
Once measurable tumors were established, bacteria were injected intravenously via the tail vein with varying dosage. Tumor-localized, bacterial gene expression was monitored in real time over the course of 60 hours using an in vivo imaging system (IVIS). At each time point, tumors were excised, homogenized, and plated to quantitate bacterial colonies for correlation with gene expression data. 
Together, this data yields a quantitative measure of the in vivo growth and gene expression dynamics of bacteria growing inside tumors.

Measuring Growth and Gene Expression Dynamics of Tumor-Targeted S. Typhimurium Bacteria

The goal of these experiments is to generate quantitative time-course data on the growth and gene expression dynamics of attenuated S. typhimurium bacterial colonies growing inside tumors. This video covers tumor cell preparation and implantation, bacteria preparation and injection, whole-animal luminescence imaging, tumor excision, and bacterial colony counting.

肿瘤靶向性生长和基因表达的动力学测量<em&gt; S。鼠伤寒沙门氏菌</em&gt;细菌

The goal of these experiments is to generate quantitative time-course data on the growth and gene expression dynamics of attenuated S. typhimurium ...

Forward Genetic Approaches in Chlamydia trachomatis

Chlamydia trachomatis, the etiological agent of sexually transmitted diseases and ocular infections, remains poorly characterized due to its intractability to experimental transformation with recombinant DNA. We developed an approach to perform genetic analysis in C. trachomatis&#160;despite the lack of molecular genetic tools. Our method involves: i.) chemical mutagenesis to rapidly generate comprehensive libraries of genetically-defined mutants with distinct phenotypes; ii.) whole-genome sequencing (WGS) to map the underlying genetic lesions and to find associations between mutated gene(s) and a common phenotype; iii.) generation of recombinant strains through co-infection of mammalian cells with mutant and wild type bacteria. Accordingly, we were able to establish causal relationships between genotypes and phenotypes. The coupling of chemically-induced gene variation and WGS to establish correlative genotype&#8211;phenotype associations should be broadly applicable to the large list of medically and environmentally important microorganisms currently intractable to genetic analysis.

Forward Genetic Approaches in Chlamydia trachomatis

We describe a methodology to perform genetic analysis in Chlamydia based on chemical mutagenesis and whole genome sequencing. In addition, a system for DNA exchange within infected cells is described that can be used for genetic mapping. This method may be broadly applicable to microbial systems lacking transformation systems and molecular genetic tools.

正向遗传途径<em&gt;沙眼衣原体</em

Chlamydia trachomatis, the etiological agent of sexually transmitted diseases and ocular infections, remains poorly characterized due to its ...

Monitoring Changes in Membrane Polarity, Membrane Integrity, and Intracellular Ion Concentrations in Streptococcus pneumoniae Using Fluorescent Dyes

Membrane depolarization and ion fluxes are events that have been studied extensively in biological systems due to their ability to profoundly impact cellular functions, including energetics and signal transductions. While both fluorescent and electrophysiological methods, including electrode usage and patch-clamping, have been well developed for measuring these events in eukaryotic cells, methodology for measuring similar events in microorganisms have proven more challenging to develop given their small size in combination with the more complex outer surface of bacteria shielding the membrane. During our studies of death-initiation in Streptococcus pneumoniae (pneumococcus), we wanted to elucidate the role of membrane events, including changes in polarity, integrity, and intracellular ion concentrations. Searching the literature, we found that very few studies exist. Other investigators had monitored radioisotope uptake or equilibrium to measure ion fluxes and membrane potential and a limited number of studies, mostly in Gram-negative organisms, had seen some success using carbocyanine or oxonol fluorescent dyes to measure membrane potential, or loading bacteria with cell-permeant acetoxymethyl (AM) ester versions of ion-sensitive fluorescent indicator dyes. We therefore established and optimized protocols for measuring membrane potential, rupture, and ion-transport in the Gram-positive organism S. pneumoniae. We developed protocols using the bis-oxonol dye DiBAC4(3) and the cell-impermeant dye propidium iodide to measure membrane depolarization and rupture, respectively, as well as methods to optimally load the pneumococci with the AM esters of the ratiometric dyes Fura-2, PBFI, and BCECF to detect changes in intracellular concentrations of Ca2+, K+, and H+, respectively, using a fluorescence-detection plate reader. These protocols are the first of their kind for the pneumococcus and the majority of these dyes have not been used in any other bacterial species. Though our protocols have been optimized for S. pneumoniae, we believe these approaches should form an excellent starting-point for similar studies in other bacterial species.

Monitoring Changes in Membrane Polarity, Membrane Integrity, and Intracellular Ion Concentrations in Streptococcus pneumoniae Using Fluorescent Dyes

Unlike that seen for eukaryotes, there is a paucity of studies that detail membrane depolarization and ion concentration changes in bacteria, primarily as their small size makes conventional methods of measurement difficult. Here, we detail protocols for monitoring such events in the significant Gram-positive pathogen Streptococcus pneumoniae utilizing fluorescence techniques.

监测改变膜极性，膜完整性和细胞内离子浓度<em&gt;肺炎链球菌</em&gt;使用荧光染料

Membrane depolarization and ion fluxes are events that have been studied extensively in biological systems due to their ability to profoundly impact ...

Using Fluorescent Proteins to Monitor Glycosome Dynamics in the African Trypanosome

Trypanosoma brucei is a kinetoplastid parasite that causes human African trypanosomiasis (HAT), or sleeping sickness, and a wasting disease, nagana, in cattle1. The parasite alternates between the bloodstream of the mammalian host and the tsetse fly vector. The composition of many cellular organelles changes in response to these different extracellular conditions2-5.
Glycosomes are highly specialized peroxisomes in which many of the enzymes involved in glycolysis are compartmentalized. Glycosome composition changes in a developmental and environmentally regulated manner4-11. Currently, the most common techniques used to study glycosome dynamics are electron and fluorescence microscopy; techniques that are expensive, time and labor intensive, and not easily adapted to high throughput analyses.
To overcome these limitations, a fluorescent-glycosome reporter system in which enhanced yellow fluorescent protein (eYFP) is fused to a peroxisome targeting sequence (PTS2), which directs the fusion protein to glycosomes12, has been established. Upon import of the PTS2eYFP fusion protein, glycosomes become fluorescent. Organelle degradation and recycling results in the loss of fluorescence that can be measured by flow cytometry. Large numbers of cells (5,000 cells/sec) can be analyzed in real-time without extensive sample preparation such as fixation and mounting. This method offers a rapid way of detecting changes in organelle composition in response to fluctuating environmental conditions.

Glycosome dynamics in African trypanosomes are difficult to study by traditional cell biology techniques such as electron and fluorescence microscopy. As a means of observing dynamic organelle behavior, a fluorescent-organelle reporter system has been used in conjunction with flow cytometry to monitor real-time glycosome dynamics in live parasites.

利用荧光蛋白监视非洲锥虫Glycosome动态

Trypanosoma brucei is a kinetoplastid parasite that causes human African trypanosomiasis (HAT), or sleeping sickness, and a wasting disease, nagana, in ...

"Phagosome Closure Assay" to Visualize Phagosome Formation in Three Dimensions Using Total Internal Reflection Fluorescent Microscopy (TIRFM)

Phagocytosis is a mechanism used by specialized cells to internalize and eliminate microorganisms or cellular debris. It relies on profound rearrangements of the actin cytoskeleton that is the driving force for plasma membrane extension around the particle. In addition, efficient engulfment of large material relies on focal exocytosis of intracellular compartments. This process is highly dynamic and numerous molecular players have been described to have a role during phagocytic cup formation. The precise regulation in time and space of all of these molecules, however, remains elusive. In addition, the last step of phagosome closure has been very difficult to observe because inhibition by RNA interference or dominant negative mutants often results in stalled phagocytic cup formation. 
We have set up a dedicated experimental approach using total internal reflection fluorescence microscopy (TIRFM) combined with epifluorescence to monitor step by step the extension of pseudopods and their tips in a phagosome growing around a particle loosely bound to a coverslip. This method allows us to observe, with high resolution the very tips of the pseudopods and their fusion during closure of the phagosome in living cells for two different fluorescently tagged proteins at the same time.

&quot;Phagosome Closure Assay&quot; to Visualize Phagosome Formation in Three Dimensions Using Total Internal Reflection Fluorescent Microscopy TIRFM

We describe an experimental setup to visualize with unprecedented high resolution phagosome formation and closure in three dimensions in living macrophages, using total internal reflection fluorescence microscopy. It allows monitoring of the base of the phagocytic cup, the extending pseudopods, as well as the precise site of phagosome scission.

“吞噬体封闭试验”可视化吞噬体形成的三个维度利用全内反射荧光显微镜（TIRFM）

Phagocytosis is a mechanism used by specialized cells to internalize and eliminate microorganisms or cellular debris. It relies on profound rearrangements ...

Precision-cut Mouse Lung Slices to Visualize Live Pulmonary Dendritic Cells

Inhalation of allergens and pathogens elicits multiple changes in a variety of immune cell types in the lung. Flow cytometry is a powerful technique for quantitative analysis of cell surface proteins on immune cells, but it provides no information on the localization and migration patterns of these cells within the lung. Similarly,&#160;chemotaxis assays can be performed to study the potential of cells to respond to chemotactic factors in vitro, but these assays do not reproduce the complex environment of the intact lung. In contrast to these aforementioned techniques, the location of individual cell types within the lung can be readily visualized by generating Precision-cut Lung Slices (PCLS), staining them with commercially available, fluorescently tagged antibodies, and visualizing the sections by confocal microscopy. PCLS can be used for both live and fixed lung tissue, and the slices can encompass areas as large as a cross section of an entire lobe. We have used this protocol to successfully visualize the location of a wide variety of cell types in the lung, including distinct types of dendritic cells, macrophages, neutrophils, T cells and B cells, as well as structural cells such as lymphatic, endothelial, and epithelial cells. The ability to visualize cellular interactions, such as those between dendritic cells and T cells, in live, three-dimensional lung tissue, can reveal how cells move within the lung and interact with one another at steady state and during inflammation. Thus, when used in combination with other procedures, such as flow cytometry and quantitative PCR, PCLS can contribute to a comprehensive understanding of cellular events that underlie allergic and inflammatory diseases of the lung.

We describe a method for generating Precision-cut Lung Slices (PCLS) and immunostaining them to visualize the localization of various immune cell types in the lung. Our protocol can be extended to visualize the location and function of many different cell types under a variety of conditions.

精密切割的小鼠肺切片可视化现场肺树突状细胞

Inhalation of allergens and pathogens elicits multiple changes in a variety of immune cell types in the lung. Flow cytometry is a powerful technique for ...

Studying Organelle Dynamics in B Cells During Immune Synapse Formation

Recognition of surface-tethered antigens&#160;by the B cell receptor (BCR)&#160;triggers the formation of an immune synapse (IS), where both signaling&#160;and antigen uptake are coordinated. IS formation involves dynamic actin remodeling accompanied by the polarized recruitment to the synaptic membrane of the centrosome and associated intracellular organelles such as lysosomes and the Golgi apparatus. Initial stages of actin remodeling allow B cells to increase their cell surface and maximize the quantity of antigen-BCR complexes gathered at the synapse. Under certain conditions, when B cells recognize antigens associated to rigid surfaces, this process is coupled to the local recruitment and secretion of lysosomes, which can facilitate antigen extraction. Uptaken antigens&#160;are internalized into specialized endo-lysosome compartments for processing into peptides, which are loaded onto major histocompatibility complex II (MHC-II) molecules for further presentation to T helper cells. Therefore, studying organelle dynamics associated with the formation of an IS is crucial to understanding how B cells are activated. In the present article we will discuss both imaging and a biochemical technique used to study changes in intracellular organelle positioning and cytoskeleton rearrangements that are associated with the formation of an IS in B cells.

Herein we describe two approaches to characterize cell polarization events in B lymphocytes during the formation of an IS. The first, involves quantification of organelle recruitment and cytoskeleton rearrangements at the synaptic membrane. The second is a biochemical approach, to characterize changes in composition of the centrosome, which undergoes polarization to the immune synapse.