Bioengineering Core for Cellular and Tissue Models

对于一张256×256像素的图像，计算完整位移图所需的时间已缩短至不到 10秒。导致该位移的最可能牵引力是 通过构建二维布西内斯克方程矩阵得出的，该方程描述了 在纯弹性半空间中，作用于表面上的剪切力 在限定区域内产生的各点位移（Butler 等，2002；Dembo 和 Wang，1999； Marganski 等，2003）。通过将 牵引节点限制在静止细胞轮廓内部的位置，生成了一个超定系统，其中 已知的位移向量数量多于牵引点位置。通过采用蒂霍诺夫正则化方法， 该方法在位移拟合到牵引大小和方向的最小二乘误差与力 平衡、扭矩平衡以及对高幅值的惩罚，生成了一个牵引图，该图最大限度地减小了 由于标记物在凝胶中随机分布不均匀而导致的位移图中的 噪声和不均匀性影响。 C3.1.f. 胶原蛋白凝胶制备。胶原蛋白和中和溶液（100 mM Hepes，pH 7.3，溶于2×PBS） 在4°C下保存，待用。 分离成年心肌细胞，并使其在 15 ml 离心管底部通过重力沉降聚集成团。将细胞稀释至每毫升 100 万至 10 万个。胶原蛋白凝胶的最终浓度为 2 mg/ml。 在冰上操作，向装有细胞的试管中依次加入培养基、中和溶液，最后加入胶原蛋白。 最终溶液很快变得粘稠，并呈现黄粉色。每块拉伸或牵引力板吸取8毫升 溶液，并将板放入37°C的培养箱中。 胶原蛋白在30-60分钟内 在细胞周围聚合，随后在凝胶上方小心加入5毫升培养基。 每3- 214 Knowlton, Kirk U. 4天更换一次培养基。对于在3D胶原蛋白凝胶中培养的成年心肌细胞，进行免疫印迹实验时，使用不含钙和镁的PBS中含0.5 mM EDTA的溶液将细胞剥离，并移除培养基。 接下来向 凝胶中加入1毫升2×裂解缓冲液，使细胞裂解，并将裂解液在4℃下离心12分钟。向60微升上清液中加入30微升3× Laemmli样品缓冲液， 在100℃下加热3分钟C下加热3分钟， 然后加载到SDS-PAGE凝胶中进行蛋白质 分析。 新方法开发：我们将把这些牵引力显微镜方法（如 C2.1.c节所示）从新生大鼠心室肌细胞扩展至新生和小鼠成年心室 肌细胞。 鉴于此前已成功对小鼠新生儿心室肌细胞进行贴壁培养和拉伸（Knoll等， 2002），我们预计这些牵引力方法无需重大 修改即可应用于小鼠新生儿心室肌细胞。然而，对于成年心室肌细胞，我们还需要在聚丙烯酰胺基质上方添加一层含有成年 心肌细胞的胶原凝胶层，以实现足够的细胞-基质粘附。鉴于我们 已观察到，分离的成年心肌细胞可在其末端通过聚赖氨酸粘附，并在不脱离的情况下产生生理性收缩 张力而不脱落（Bluhm 等，1995），且我们还观察到新生小鼠心肌细胞在 附着于聚丙烯酰胺凝胶时能够产生生理性活性应力（C2.1 节C），我们确信 能够创造出使成年小鼠心室心肌细胞充分粘附于 足够凝胶的条件，从而产生可测量的基质变形，进而计算牵引力。细胞将 嵌入含有聚赖氨酸的胶原蛋白凝胶中，以增强细胞附着强度。 胶原蛋白凝胶将通过 将1.55 mg/ml的I型胶原蛋白（BD Biosciences）混入pH 7.3的100 mM HEPES缓冲液中，并将 该溶液置于37°C下30分钟使其凝胶化而制备。 还将制备另一种凝胶溶液，其中含有相同浓度 的胶原蛋白（1.55 mg/ml），溶于含BDM的心脏缓冲液中，并加入2.5%直径为500 nm的荧光素 包被聚苯乙烯微球（Polysciences）、0.05 mg/l聚-L-赖氨酸-琥珀酰亚胺（Sigma），以及基于凝胶表面积计算的 10,000 个细胞/cm² 的心肌细胞 悬液。将该溶液用移液管滴加在 原始溶液表面，室温静置30分钟后，置于37°C环境中30分钟。 这种低温凝胶化处理可使细胞和微珠沿同一平面沉降至下表面，从而 便于显微镜观察和分析。 凝胶聚合后，用含2 mM钙离子的泰罗德 溶液冲洗3次，并让其浸泡在该溶液中，直至细胞开始收缩。 C3.1.g. 心肌细胞微图案化。 我们开发了两种对可变形 膜进行微图案化的方法，用于构建图案化心肌细胞。这些方案结合了光刻、微流控和 微沟槽技术，在弹性膜上微图案化细胞粘附位点，并结合心肌细胞 及成纤维细胞的分离与培养相结合。由此建立的心肌组织体外模型模拟了原生心室心肌的重要结构 和功能特征，在更好地保持细胞形态、 排列和连接性的同时，还能对细胞-基质粘附和机械环境（例如 施加拉伸力）进行调控。 这两种方法均在《自然》 Protocols上即将发表的一篇新文章中进行了详细描述（Camelliti等人，2006）。最初的微流控图案化技术（也称为微接触 印刷）使用平坦的硅橡胶膜，在其表面沉积由细胞外 基质蛋白组成的条纹以引导细胞附着（Gopalan等人， 2003）。该技术如图12A所示， 涉及一个可重复使用的微结构硅橡胶印章，该印章由光刻蚀制的 电子级硅金属晶圆浇铸而成。 将印章手动压印在目标膜上，形成 微流体通道，用于在空间上受限的区域施加基质蛋白溶液。溶液干燥后， 小心地移除印章，在膜上留下具有图案化的生长基质。 可以 进行双重印制，例如创建平行线规则交错的图案，从而 将伪ID结构（培养细胞的线状排列）转化为组织的伪二维表示（网格）。 已成功将分离的心肌细胞接种在由胶原蛋白制成的此类轨道上。 该方法 具有良好的空间分辨率和空间各向异性表现能力，但会导致细胞培养密度较低。 通过从 蚀刻硅片上制备聚二甲基硅氧烷（PDMS）复制品，可实现深层微结构的微加工。 随后利用该模具，通过微流控技术制备胶原蛋白、层粘连蛋白、纤维连接蛋白或其混合物的 图案。5分钟后，移除微流控模具并洗去多余的基质蛋白， 使图案保留在基底上。随后，如上所述，将心肌细胞培养在膜或培养皿上。 在图案化基底上，细胞会附着、铺展、生长，并呈现出细长的类组织表型。 由于在拉伸操作过程中细胞处于血清饥饿状态（以防止非拉伸诱导的细胞增殖）， 血清中蛋白质的非特异性吸附极少。远离 215 Knowlton, Kirk U. 基底图案区域之外的细胞粘附，也可通过用聚乙二醇 聚合物（Li 等，1996）或共价结合的聚乙二醇（PEG）修饰疏水表面来减少。 负显影 光刻胶 透过掩模接受紫外线照射 ¿透明掩模 ^显影曝光后的光刻胶 微图案化 SI晶圆 [浇铸POMS模具 带 微通道的PDMS主体 膜上的PDMS模具 用胶原蛋白填充通道 在培养箱中孵育h 去除PDMS模具 膜上的胶原蛋白 轨道 细胞位于平行的 胶原蛋白轨道上 膜上的POMS模具 '用胶原蛋白填充通道 1 h 3T*C 培养箱中 1 去除PDMS模具 40'C 过夜 膜上交叉的胶原蛋白 细胞在交叉胶原蛋白轨道上 交叉胶原蛋白轨道上的细胞 胶原蛋白轨迹上 图12. A. 微图案化流程。B. 等双轴和非等双轴细胞拉伸器的组装 第二种微图案化方法是在硅胶膜本身上制作沟槽（图13）。 采用此方法，细胞在膜上的汇合度更高，因此每个拉伸器可提供更多的蛋白质供分析 。硅片的光刻图案化按照先前描述的方法进行 （Bhatia 等，1998）。简而言之， 在硅片上涂覆负性光刻胶，通过透明掩模曝光于紫外线（365 nm）下，并采用高分辨率印刷工艺显影，以形成具有所需宽度 和平行线间距（10 µm平行线及10 µm间距）的图案。聚二甲基硅氧烷 （PDMS）由两种液体组分混合制备（Sylgard 186 试剂盒，道康宁公司），作为薄层浇注到 显影后的晶圆上，经脱气处理后固化。所得膜层涂覆有 用于对齐心肌细胞的所需细胞外基质（ECM）。 图13. 用于 膜微图案化 的新型心肌细胞图案化实验，可改善因慢性细胞拉伸而引起的 蛋白质变化的定量测量。先制备一个PDMS PDMS模具，并利用该模具制作具有相同对准 图案的薄 硅胶膜。 在表面涂覆一层 硅胶膜 胶原蛋白（或其他基质），并将心肌细胞接种到 膜上。细胞会沿膜 对齐并呈现细长、棒状的 胶原涂层表型，随后对这些 细胞制备物施加拉伸模式（纵向 和/或横向）。 心肌细胞 沿图案排列 C3.1.h. 拉伸装置与膜。采用与 Sadoshima 等（Sadoshima et al., 1992）类似的方法，制备了涂覆 I 型胶原蛋白及其他基质的 Silastic 可变形膜。 将细胞（2 × 10⁵ 个/cm²）接种在涂覆有I型胶原蛋白的带图案或无图案硅膜上，并安装 在等双轴（均匀）或非等双轴（椭圆形）拉伸装置上，如图12B所示 （Lee等人，1996）。 细胞在无血清条件下培养24小时，随后进行24小时的被动拉伸。提取总RNA ，并使用Direct RPA系统（Ambion Inc.）进行RNA保护实验（RPA）。小鼠BMP 216 Knowlton, Kirk U. 编码序列通过RT-PCR扩增，并作为BMP核糖探针合成的模板，同时以 小鼠GAPDH作为对照。 微图案化技术使对齐的心肌细胞培养物能够在椭圆形拉伸器上接受非 等轴向拉伸（Gopalan 等，2003），该拉伸器以恒定比例（例如 2:1） 向细胞施加各向异性应变。若细胞排列时其轴线与椭圆形拉伸仪的一个轴线对齐，则 主应变比由椭圆的小径与大径之比决定。 借助这些设备， 应变的大小可在任意值下精确控制，且每台拉伸仪均经过校准。 C3.1.I. 组织学细胞制备。 为了评估在 不同刚度凝胶上培养的心肌细胞中肌节的清晰度与组织结构，在初始接种后48 小时和7天时对细胞进行固定并染色，以检测a-肌动蛋白等标志物。 细胞固定方法为：先用冷（4°C）PBS洗涤，随即 浸入冷0.75%戊二醛溶液中20分钟。 用0.1% Triton X100（Sigma）处理5分钟使细胞膜通透。抗体用1%牛血清白蛋白 （BSA，Gemini）中稀释，并加入 至固定好的细胞中，在室温下孵育60分钟。四荧光素异硫氰酸酯 （TRITC）标记的山羊抗兔二抗（Jackson Immunoresearch，宾夕法尼亚州西格罗夫）用1% BSA稀释后， 加入并孵育30分钟。随后使用NCMIR的共聚焦显微镜，或通过相位和 落射荧光模式下进行成像，并使用Metamorph软件（Universal Imaging Corporation）采集图像。 此外，通过使用ImageJ软件（NIH）对细胞进行分割，计算细胞面积和 纵横比（长轴/短轴）。 C3.1.I. 细胞内钙瞬变的比值成像。将分离的细胞用Fura-2 进行标记（Grynkiewicz 等，1985）。 Fura-2钙指示剂（Molecular Probes）在 高钙泰罗德缓冲液（130 mM NaCl、5.4 mM KCl、2 mM CaCl、1 mM MgCl₂、0.3 mM Na₂HPO₄、 10 mM HEPES、5.5 mM 葡萄糖，pH 7.4）中稀释至5 μM浓度，并在室温下孵育30分钟。随后用新鲜的泰罗德缓冲液洗涤细胞3 次，并在整个实验过程中将细胞置于泰罗德缓冲液中。细胞 使用尼康Eclipse TE3000倒置显微镜进行成像，并按照与牵引力测量相同的程序 刺激其搏动。拍摄静息心肌细胞的相差图像，以便 限定图像分析范围。使用Lambda DG-4高速滤光片更换器 （Sutler Instruments），细胞 交替接受380 nm和340 nm光照，并使用 Photometries Cascade 51 2X CCD相机对510 nm处的荧光进行成像，数据通过Metamorph软件 （Universal Imaging Corporation）进行采集。每15毫秒切换一次滤光片并采集新图像，持续4.5秒（共300张图像）。 为了校准Fura-2的响应，加入等量的高钙泰罗德缓冲液，其中含有20 µM 钙离子载体-4-溴-A23187（Molecular Probes）的高钙泰罗德缓冲液，以饱和结合在钙上的Fura， 并在340 nm和380 nm激发/510 nm发射波长下采集图像。 移除培养液，并加入 含有10%乙二胺四乙酸（EDTA，Sigma）的培养液，以螯合所有钙离子， 从而获得Fura的基线荧光信号。再次在340 nm和380 nm激发/ 510 nm发射波长下再次采集图像。 使用在Metamorph或ImageJ（NIH）中编写的自定义宏对图像进行分析，将 交替采集的340 nm和380 nm激发波长图像序列拆分为两个图像堆栈，计算这两个堆栈的比值 并计算细胞轮廓内每张图像该比值的平均值（针对手动勾勒的细胞 区域）。随后，利用以下公式计算细胞内游离钙的浓度： -R] max / 其中 kd 为解离常数（0.14 μM），Q 为 380 nm 激发下细胞轮廓内总平均荧光强度之比， R 为每张图像中 340 nm 激发荧光强度与 380 nm 激发荧光强度之比 激发下各图像中荧光强度的比值，Rmin 是添加 EDTA 后的该比值，f?ma* 是添加 钙离子载体后的该比值。 C3.1.k. 细胞内信号传导事件的FRET成像。将新生儿心肌细胞转染 重组FRET报告基因，例如A-激酶活性报告基因AKAR2（Zhang等，2005；Zhang等， 2001），并在转染后2天进行成像（转染效率为-5%）。 C3.1.k. 细胞内信号事件的FRET成像。将新生儿心肌细胞在分离后1天使用FuGeneS转染试剂（罗氏诊断）转染 重组FRET报告基因（例如A-激酶活性报告基因AKAR2（Zhang等，2005；Zhang等， 对于成年细胞，我们将采用病毒介导的基因转移。AKAR2是一种 由GFP的黄色（YFP）和青色（CFP）突变体、蛋白激酶A 底物，以及一个磷酸苏氨酸结合域。PKA介导的AKAR2磷酸化会增加FRET 217 Knowlton, Kirk U. ，从而可通过黄/青 荧光发射比值对内源性PKA活性进行实时荧光成像。表达报告基因的固定心肌细胞经a-肌动蛋白标记，以确认细胞形态正常。 在成像前20分钟，用含20 mM HEPES缓冲液（pH 7.2）的汉克斯缓冲盐溶液洗涤细胞， 并在成像过程中使用载物台加热器（Carel）将温度维持在35°C。转染的心肌细胞 在配备60X Plan Apochromat物镜的尼康Eclipse TE300倒置显微镜上进行成像， Photometries Cascade 512F CCD相机及MetaFluor 6.2软件（Universal Imaging Corporation）进行成像。 成像时使用430/25 nm激发滤光片（用于CFP），并同时记录CFP（470/30 nm）和YFP发射（535/30 nm）信号。 每5秒采集一次1秒曝光的图像。为获得荧光强度比的时间曲线，将每张图像中的YFP或CFP荧光 强度在感兴趣区域内进行平均，扣除背景，然后计算黄色/青色 荧光比值，并以试剂添加前的比值进行归一化。对于荧光比值图像， 采用类似方法逐像素处理，并使用MetaMorph软件 （Universal Imaging Corporation）进行5×5像素中值滤波。 C3.2. 分离的小鼠小梁和乳头肌 C3.2.a. 心肌分离与固定。用异氟烷麻醉雄性或雌性小鼠。颈椎 脱位后，打开胸腔，并通过心内注射低钙、高 钾心脏停搏液使心脏停搏。 随后迅速取出心脏，插入导管，并用改良的 Hepes缓冲液进行灌注，该溶液含（mM）：137.2 NaCI、15.0 KCI、1.2 MgCI2、2.8 mM 乙酸钠、10 牛磺酸、 1.0 CaCl₂、10 mM 葡萄糖及 10 mM Hepes，该溶液与 100% 氧气处于平衡状态。切开右心室， 通过先切除一小段瓣膜（该瓣膜通过腱索与 肌纤维通过腱索附着于瓣膜，随后切开一小段室间隔壁——该肌纤维 在基部附着于此。乳头肌的选择基于其几何形状。通常，此类研究中选用的乳头 肌应为细长且无分叉的。 研究中不采用左心室（LV） 乳头肌，因为这些肌体较厚，且氧气和营养物质向其核心部位的 扩散可能受到限制。 此外，左心室乳头肌的氧气供应减少可能影响ANP和BMP的诱导， 因此这些标本可能无法产生正常的牵张诱发反应（Chen, 2005；Sabatine等，2004）。 将肌肉组织安装在心脏组织培养腔中，该腔内盛有与解剖时相同的Hepes缓冲溶液。 肌肉组织安装在连接力传感器和由执行器 控制的微操纵器的篮筐，以及一个类似钛钩的固定延伸件之间。在安装过程中，氧气会流过溶液 ，以防止缺氧。对于持续时间超过6 小时的实验，系统所有部件在每次实验前均需进行蒸汽灭菌。 此外，所有溶液均经滤器灭菌。随后， 将Hepes缓冲液更换为改良的M199细胞培养基，其成分（mM）为：2.0 L- 肉碱、5.0 肌酸、5.0 牛磺酸、 2.0 L-谷氨酰胺、0.2% 白蛋白、100 IU/ml 青霉素、0.1 mg/ml 链霉素、10 mM Hepes 以及 50 ug/ml 胰岛素，并在 5% CO₂ 和 95% O₂ 的平衡环境中进行。肌肉腔室的 温度维持在34°C。溶液的pH值维持在7.4至7.5之间。 将改良的Hepes缓冲溶液分步替换为改良的M199培养基，使钙 浓度从1.0 mM缓慢升至1.75 mM。溶液更换完成后，通过钩子及置于肌肉基部附近的铂电极对肌肉进行 刺激。 将肌肉保持在松弛长度，并以0.2 Hz的频率刺激一小时（Janssen等，1998）。待肌肉 达到平衡状态后，将其拉伸至Lmax的85-95%（其中Lmax定义为 肌肉产生最大主动力的长度），或保持在松弛长度状态直至 实验结束（2小时、5小时、12小时）。 肌肉长度通过与微操纵器并联的LVDT进行测量。 可在每块肌肉表面用二氧化钛标记物进行标记，以便在实验过程中 观察局部肌肉变形。肌肉尺寸通过视频采集获得， 并利用放置在标本焦平面上的已知尺寸模拟体进行校准。 C3.2.b. 力学测试。 右心室乳头肌安装在固定钩与 哈佛仪器等长力传感器（724490）之间，该传感器可测量 0-0.5g 范围内的力，精度为 ¿1%（<1 mg）。哈佛仪器力传感器连接至 Newport 460P 系列高精度线性模块化滚珠轴承载物台，其在 x/y/z 方向的位置由 微操纵器控制，该微操纵器连接至 Newport 电动执行器 （CM-12CC）相连，该执行器的 分辨率小于100 nm，增量位移小于500 nm，速度范围为50-500 218 Knowlton, Kirk U. Urn/秒。为了以 约1 |xm的精度测量肌肉长度及肌肉拉伸距离，将一个LVDT (Omega LD310-10) 与执行器并联安装，该LVDT的精度为400 nm，线性量程为20 mm， 。 利用该实验装置，在以1 Hz频率刺激肌肉以及 肌肉连续拉伸至Lmax的过程中，记录单轴肌肉力。同时采集肌肉长度，并由CCD相机（COHU Inc.） 成像肌肉的局部变形。肌肉 变形的数字视频记录与采集到的力数据保持同步。基于每条肌肉的松弛长度，计算拉格朗日单轴应变（E=1/2(A²-1)， 其中A为拉伸比）。拉格朗日应力 则通过将采集到的力数据除以每条肌肉的初始横截面积来计算。 C3.2.C. 实时PCR操作流程。每次肌肉实验结束后，立即将肌肉 浸入RNA Later溶液中，该溶液可稳定完整组织样本中的mRNA，并含有 可灭活RNase的试剂（Qiagen）。将右心室乳头肌完全从其 附着处、瓣膜及室间隔壁中分离出来。随后对肌肉进行匀浆处理，并使用Qiagen的RNeasy Micro RNA纯化试剂盒从这些 组织中提取RNA。鉴于从这些微小 样本中总RNA的含量较低（-10 ng/μL），且已有研究表明DNase处理会降解和/或破坏核酸 提取物，因此未对右心室乳头肌样本进行DNase处理（Jemiolo和Trappe，2004）。 反转录（RT）使用Invitrogen SuperScript III cDNA合成试剂盒完成，该试剂盒 针对低输入量的RNA进行了优化。最后，使用Applied Biosystems ABI 7700 实时热循环仪，使用含UNG的AB TaqMan通用主混合液，以及针对ANP、BMP和GAPDH的AB预制引物和TaqMan 探针。 在离体心肌细胞以及完整心肌组织中，ANP和BNP基因的表达通常会因机械负荷增加而 被诱导（Ruwhof等，2000）。GAPDH管家基因的表达在这些实验中 保持不变。对于每份测试的RNA样本，均获得两组Ct值并 计算其平均值。 随后对所得的平均Ct值应用AACt方法，以定量测定ANP或BNP相对于未拉伸对照样本的 倍数变化。在每个基因表达检测中， 均包含相应的对照。对于每份RNA样本，均未进行无RT对照，以检测是否存在 基因组DNA污染。 此外，还进行了阴性对照（即未向反应体系中添加cDNA）， 以检验酶主混合液以及引物和探针混合物的纯度。数据 以均值 ± 1SEM 形式呈现。 由于mRNA浓度呈对数正态分布且需要 采用非参数统计方法，因此对拉伸组与未拉伸实验组之间的ACt值或从 热循环仪获得的原始输出数据，进行了两组t检验。显著性水平设定为P < 0.05，并对多重比较进行适当校正。 C3.2.d. 抗体染色与共聚焦显微镜观察。 未用于基因表达 分析的乳头肌被拉伸至已知的拉伸百分比，并用2% PFA（对甲醛）固定，该溶液由50% PBS （磷酸盐缓冲盐水）和50%改良HEPES缓冲心肌停搏液稀释而成。 10-20分钟后， 将溶液更换为100% PBS中的4% PFA。让肌组织继续固定10-20分钟， 然后转移至Tissue-Tek O.C.T.化合物中。将其置于干冰上冷冻，并储存在-80¿C。随后使用 冷冻切片机对组织样本进行切片（10-15 u,m 厚切片）。对于ANP蛋白分析，制备组织横 切片；而对于巨蛋白表位染色，则制备纵切片。经过一系列 封闭和洗涤步骤后，组织切片用相应的抗体进行染色 （例如，使用IgG抗小鼠 ANP抗体检测拉伸和未拉伸标本中的ANP蛋白表达；IgG 单克隆抗小鼠α-肌动蛋白（Sigma）抗体用于染色肌节Z盘，单克隆抗小鼠IgM 9D10抗体用于染色巨蛋白上的PEVK区域（Hybridoma Bank）， 以及多克隆抗山羊IgG特莱托宁 抗体用于染色Tcap（Santa Cruz Biotechnology），该蛋白与巨蛋白的N端相连）。 还使用了与Alexa 568或Alexa 488偶联的相应二抗。 对于巨蛋白表位 染色程序，将组织分别与α-肌动蛋白和Tcap或α-肌动蛋白和9D10进行双重染色。使用 Biorad共聚焦显微镜，配合适于同时激发Alexa- 568和Alexa-488的滤光片组对切片进行成像。 巨蛋白表位与Z盘之间的距离是利用 FFT分析测定的，这与我们先前针对骨骼肌纤维的报道一致 （Shah 等，2004）。 219 Knowlton, Kirk U. C3.3. 离体灌注小鼠整体心脏制备 C3.3.a. 离体小鼠心脏制备。向小鼠腹腔注射肝素（100 USP单位），用异氟烷麻醉，并通过颈椎脱位处死。迅速摘除心脏， 用冷心肌停搏液冲洗，并将主动脉插入20号不锈钢导管中。 将 主动脉周围的剩余组织用5-0缝线扎紧，以阻断心脏与所有血管的连接。将 心脏安装在逆行兰根多夫灌注装置（Radnoti）上。 冠状动脉灌注液为 含氧的改良克雷布斯-亨塞莱特溶液：24.9 mM NaHCO₃、1.2 mM KHaPO₄、11.1 mM 葡萄糖、1.2 mM MgSO₄、4.7 mM KCl、118 mM NaCl 以及 2.55 mM CaCl₂（Grupp 等，1993）。 灌注液中通入 95% O₂–5% CO₂ 混合气体，温度维持在 35–37°C，并通过施加 70 mmHg 的恒定压力使流速达到 1–2 ml/min。清除冠状动脉中的残留物后，让心脏自由搏动并 平衡15分钟。放置表面心电图（EGG）电极，并在整个 实验过程中进行记录。 95% O2 / 5% CO2 采用数字起搏器（DS8000，World Precision Instruments）对心脏进行起搏。 通过双极铂电极从心房或心室进行 逆行 灌注，以恒定电流（为阈值的两倍）进行心外膜起搏。 储液囊 对于等容 兰根多夫制备，将充液气球经二尖瓣插入 顺行左心室。 该气囊， 灌注 储液囊由20号钝头 针尖端的保鲜膜构成，连接到注射器输液泵上，并与 充满液体的压力传感器串联。 将气囊充气 顺应性腔 并放气至30 mmHg的末期扩张压（EDP），以对 莫安主动脉P 测量组织进行预处理。对于工作心脏制备，心脏收缩 流出道阻力 时需对抗由一个含有气泡的腔室 f. 心输出量 和一根带有可调节 夹子的柔性管组成的阻抗-顺应性网络（图14）。除拍摄标记 变形视频外，整个实验过程中心脏始终浸没在温（37°C） 收集 储液罐KHS中，除非正在对标记 变形进行录像。主动脉 右心房流出道管线中的压力通过传感器（Millar Pacing Loads Instruments，德克萨斯州休斯敦）进行监测。 摄像机 图14. VCR Core A实验室中离体射血心脏装置的 示意图。（摘自Karlon等人，2000） C3.3.b. 非均匀心外膜应变分析。 通过高分辨率光学成像技术，对孤立小鼠心脏中 50-100个二氧化钛微小标记物的分布进行映射，这些标记物使用短毛刷 排列在心外膜上。 除了左心室应变外，我们实验室还利用 这些方法绘制了离体小鼠工作心和非工作心 中室间隔应变的区域分布图（Karlon et al., 2000），并重建了收缩期右心室应变 （Lorenzen-Schmidt 等人，2005）。将使用双极刺激电极从右心房对心脏进行起搏，并使用高 保真度的米勒压力-容积导管测量心室容积和压力。 将使用一台 高分辨率CCD相机，在等容期或射血期收缩过程中，针对整个 舒张末期容积范围内的心外膜进行成像。利用MetaMorph（Universal Imaging）软件包中的目标追踪工具， 在整个心脏周期内以亚像素精度追踪标记点。 将标记点 映射到双三次赫尔姆霍兹有限元网格上，并利用标记点位移的最小二乘拟合， 通过麦卡洛克博士 实验室开发的软件计算二维非均质应变场 （Karlon 等，2000；Mazhari 等，1998）。简而言之，心外膜表面采用 6-8 个双三次 赫尔姆霍兹长椭球有限元进行建模，并从 视频图像中测得的二维参考标记坐标被投影到该表面上。 随后，变形后的坐标被投影到模型中相应的 材料点上，再通过最小二乘法进行重新拟合，从而得到一个参数化变形的 网格，据此可在每帧中插值出各区域的应变。 该方法还曾与 离体灌注兔心电活动的光学映射技术结合使用（Sung 等，2003）。除 计算等体积应变 （其数值相当大）外，还可以通过采用舒张期末帧（此时心室容积处于中高水平）作为参考状态， 以及收缩期末帧（此时心室容积较低）作为变形状态，从而获得代表 非工作心脏射血期壁运动的应变。这些 220 Knowlton, Kirk U. 这些技术均可应用于右心室和左心室，这对项目2 （Chen）至关重要。 C3.3.C. 动作电位传导的光学映射。将0.8毫升电压敏感染料di-4- ANEPPS（25 μM）的0.8毫升 bolus 注入灌注管路。心室心外膜由两盏波长为470 nm 、功率为3.6 W、含40个LED的集群灯泡（Ledtronics, Inc.）进行照射。荧光发射光通过>610 nm的高通滤光片，经一支50 mm高速镜头（1:0.95，Navitar）聚焦，并由一台12位电荷耦合 器件（CCD）相机（CA-D1-0128T，Dalsa）。以950帧 每秒的速率和64×64像素的空间分辨率采集9×9 mm²视场的图像。 荧光图像的时间序列提供了每个像素的时间强度信号。 每个信号均 以基线强度（AF/F）为基准进行归一化，并进行反相处理，以更好地反映心脏动作电位。 采用空间相位偏移和时间中位数滤波（Sung, 2001）来降低信号中的噪声。 激活时间定义为动作电位上升段中斜率（dF/dt）达到最大值的时间点。 这 可生成覆盖心室心外膜的激活图，显示动作电位 传导的速度和路径。利用激活时间的空间梯度来计算每个像素处的局部表观传导速度向量 （Bayly 等，1998）。此外，激活波前的曲率被计算为 归一化传导速度矢量场的空间梯度。 复极化时间通过 识别信号峰值，并确定动作电位分别恢复至基线水平的20%、50%和80%时 来计算。随后，动作电位持续时间（APD）被定义为复极化与 激活之间的时间差。 APD离散度定义为心脏表面APD的标准差。 在光学映射研究中，为分析复极化现象，必须对运动信号进行衰减处理。采用改良的 克雷布斯-亨塞莱特（Krebs-Henseleit）溶液，其中含有1 mM CaCb和15 mM的电机械解耦剂2,3-丁二酮 单肟（BDM），在数据采集期间用于灌注，以消除运动伪影。然而，已有研究 研究表明，BDM会延长离体小鼠 心脏的动作电位持续时间并减慢传导速度（Baker 等，2004）。 我们最近开发了一种结合比值法和运动追踪 算法的方案，可在不使用化学解耦剂的情况下消除运动伪影。由于 电位敏染料的发射光谱会随膜电位的变化而发生波长偏移，因此可以分别在绿色（560 nm）和红色 （620 nm）波长下分别记录正向和反向光学动作电位。通过 波长分离光学装置，我们能够利用单台CCD相机同时记录低波长和高波长的荧光发射， 该技术最初应用于比值显微镜（Kinosita等，1991）。 对两个波长的图像进行配准和比值计算，可放大记录到的光学 动作电位幅度，并校正因染料负载不均或 激发光不均匀导致的像素强度值差异。为校正收缩过程中心肌的运动，采用卢卡斯-金田（Lucas-Kanade）光流 算法，计算帧间像素位移的向量场，进而对变形帧进行非 刚性亚像素分辨率变换，将其恢复至参考配置。综合运用 这些技术，可有效校正光学记录中的组织位移和局部运动伪影。 C3.3.d. 程序化刺激。通过程序化S1-S2刺激方案，评估 离体小鼠心脏的复极动力学及对心律失常的易感性。使用数字刺激器（World Precision Instruments, DS8000）通过双极铂电极从左 心室心外膜对离体心脏进行起搏。 以200毫秒（S1-S1）的基本周期长度对心室进行起搏，持续>20次搏动后，随后施加 过早刺激（S2）（Baker等，2000；Lerner等，2000）。 S1-S2周期间隔逐渐缩短，直至 达到有效不应期，即S2刺激不再诱发动作电位时。在 该实验方案中，采用光学映射技术分析动作电位形态（振幅和持续时间）以及 S2诱发的动作电位的传导速度（Knollmann 等，2006）。这些 测量指标与S1-S2舒张期间隔之间的关系可用于评估小鼠 心脏的复极动力学。在S1-S2周期较短时出现的早搏可在易感 组织中诱发心律失常。可通过该实验方案中室性 心动过速的发生率来评估转基因心脏的心律失常易感性（Lerner 等，2000）。此外，成串起搏或一连串高 频 S2 刺激可在离体心脏中诱发心律失常（图 15）。 C3.3.e. 电学空间常数和时间常数。有效空间常数和时间常数直接综合了 心肌的电阻和电容，其测量方法与 Poelzing 等人的方法类似， 适用于离体小鼠心脏。 (Poelzing 等，2005)。使用一根直径为 125 微米、聚四氟乙烯涂层的铂金 221 Knowlton, Kirk U. 单极电极对左心室中外侧游离壁进行起搏。在电极进行稳态起搏 进行稳态起搏（100次起搏搏动）后，在最后一次起搏脉冲序列 刺激后35毫秒处，即前一次动作电位的不应期内，施加1 mA的阴极刺激，持续250毫秒，从而在 释放250毫秒脉冲时产生阴极中断刺激。 信号按基线动作电位振幅进行归一化处理，以便 在不同空间位置间比较组织对刺激的响应。共模信号（刺激远端处的 平均信号

the time to compute a full displacement map was reduced to less than 10 seconds for a 256x256 pixel image. The most likely traction force that can cause this displacement was found by generating a matrix of the Boussinesq equations in two dimensions, which describe the displacements at every point in a purely elastic half space that are produced by a force shear to the surface acting over a defined area (Butler et al., 2002; Dembo and Wang, 1999; Marganski et al., 2003). By limiting traction nodes to locations inside of the resting cell outline, an over-specified system was generated in which more displacement vectors were known than traction locations. By using a method of Tikhonov regularization, which balances the least squares error of a displacement fit to traction magnitudes and directions with a force balance, torque balance and penalty for high magnitudes, a traction map is generated that minimizes the effects of noise and non-uniformity of the displacement map due to non-uniformity in the random distribution of markers in the gel. C3.1.f. Collagen Gel Preparation. Collagen and neutralizing solution (100 mM Hepes, pH 7.3 in 2* PBS) are kept at 4¿C until needed. Adult cardiac myocytes are isolated and allowed to aggregate by gravity at the bottom of a 15ml tube. Cells are diluted to 1 million 100,000 per ml. Collagen gels have a final concentration of 2 mg/ml. Media, neutralizing solution and then collagen are added to the tube containing the cells working on ice. The final solution quickly becomes viscous and takes on a yellow/pinkish hue. We pipette 8 ml per stretching or traction force plate and place the plate in the incubator, at 37¿C. The collagen polymerizes around the cells in 30-60 minutes, and 5 ml media is then carefully added on top of the gel. Media is changed every 3- 214 Knowlton, Kirk U. 4 days. For immunoblotting of adult cardiac myocytes cultured in 3D collagen gels, cells are detached with 0.5 mM EDTA in calcium and magnesium-free PBS and media is removed. Next 1 ml 2* lysis buffer is added to the gel, the cells are lysed and the lysate centrifuged at 4¿C for 12 minutes. 30 ul 3* Laemmli sample buffer is added to 60 ul supernatant, heated for 3 minutes at 100¿C, and loaded into an SDS-PAGE gel for protein analysis. NEW METHOD DEVELOPMENT: We will extend these traction force microscopy methods (demonstrated in section C2.1.c) from neonatal rat ventricular myocytes to apply to them neonatal and adult mouse ventricular myocytes. Having previously successfully plated and stretched mouse neonatal ventricular myocytes (Knoll et al., 2002), we expect these traction force methods to apply to mouse neonatal myocytes without significant modification. However, for adult myocytes, we will also need to add a layer of collagen gel containing the adult myocytes on top of the polyacrylamide substrate to achieve adequate cell-matrix adhesion. Given that we have seen isolated adult myocytes can be adhered at their ends with poly-lysine and generate physiological systolic tension without detaching (Bluhm et al., 1995), and given that we have also seen that neonatal myocytes can generate physiological active stresses when adhered polyacrylamide gels (section C2.1.C), we are confident that conditions can be created that will allow adult murine ventricular myocytes to be sufficiently adhered to a sufficiently gel to generate measurable substrate deformations and thus to calculate traction forces. Cells will embedded in a collagen gel containing poly-lysine to increase cell attachment strength. The collagen gel will be formed by mixing 1.55 mg/ml collagen type I (BD Biosciences) in 100 mM HEPES buffer at pH 7.3 and placing this solution at 37¿C for 30 minutes to gel. Another gel solution will be made containing the same concentration of collagen (1.55 mg/ml) in the BDM-containing heart buffer solution plus 2.5% 500 nm diameter fluorescein- coated polystyrene beads (Polysciences), 0.05 mg/l poly-L-lysine-succinimide (sigma) and a myocyte suspension of 10,000 cells/cm2 based on the gel surface area. This solution will be pipetted on top of the original solution and allowed to sit at room temperature for 30 minutes before placing at 37¿C for 30 minutes. This low-temperature gelation allows the cells and beads to settle to the lower surface along one plane to facilitate microscopy and analysis. When the gel has polymerized, it will be washed 3 times with Tyrodes solution containing 2 mM calcium and allowed to sit in that solution until the cells begin to contract. C3.1.g. Myocvte Micropatterninci. We have developed two methods for micropatterning deformable membranes for engineering patterned myocytes. The protocols combine photolithographic, microfluidic and micro-grooving techniques, to micropattern cell adhesion sites on elastic membranes, with cardiac myocyte and fibroblast isolation and culture. The resulting in vitro models of cardiac tissue mimic important structural and functional aspects of native ventricular myocardium, and show improved preservation of cell-shape, alignment and connectivity, while providing control over cell-matrix adhesion and mechanical environment (e.g. application of stretch). Both of these approaches are described in detail in a new article in press in Nature Protocols (Camelliti et al., 2006). The original microfluidic patterning technique (also known as microcontact printing) uses flat silicone rubber membranes, onto which stripes are deposited that consist of extra-cellular matrix proteins to guide cell attachment (Gopalan et al., 2003). This technique is shown in Fig. 12A and involves a reusable microstructured silicone rubber stamp, which is cast from photolithographically etched electronics-grade silicon metal wafers. The stamp is manually sealed onto the target membrane, forming microfluidic channels for spatially-restricted application of matrix protein solution. After drying of the solution, the stamp is carefully removed, leaving a patterned growth substrate on the membrane. It is possible to double-print structures, for example to create regularly intersecting patterns of parallel lines, thereby transforming pseudo-ID structures (lines of cultured cells) into a pseudo-2D representation of tissue (grids). Isolated cardiac cells have been successful plated onto such tracks, made of collagen. This procedure provides good spatial resolution and definition of spatial anisotropy, but results in sparse cultures. Microfabrication of deep microstructures is achieved by molding a replica of polydimethylsiloxane (PDMS) from etched silicon wafers. The mold is then used to create a pattern of collagen, laminin, fibronectin, or a mixture by microfluidics. After 5 minutes, the microfluidic mold is removed and excess matrix protein washed away, leaving the pattern on the substrate. Myocytes are then cultured onto the membrane or dish as described above. On the patterned substrata, cells adhere, spread, grow and take on an elongated tissue-like phenotype. Non-specific absorption of proteins from serum is minimal since the cells are serum starved during the stretching protocols (to prevent non-stretch-induced cellular growth). Cellular adhesion away from the 215 Knowlton, Kirk U. substrate patterns can also be reduced by modifying the hydrophobic surfaces with polyethylene oxide polymers (Li et al., 1996) or covalently coupled polyethyleneglycol (PEG). Spin negative photoresist le to UVthrough mask ¿Transparency mask ^Develop exposed photoresist Mlcropatterned SI wafer [Cast POMS mold PDMS main with micro-channels PDMS mold on membrane Fill channelswith collagen h arc In Incubator Remove PDMS mold Collagen tracks on membrane Cells on parallel collagen tracks POMS mold on membrane 'Fill channels with collagen 1 h3T*C in Incubator 1 Remove PDMS mold 40'C overnight Criss-cross collagen Plate cella tracks on membrane Cells on criss-cross A collagen .tracks Figure 12. A. Micropatterning procedures. B. Equibiaxial and non-equibiaxial cell stretcher assembly The second method for micropatterning involves fabricating grooves into the silicone membrane itself (Fig. 13). With this method, cells are more confluent on the membrane, and thus more protein is available for analysis from each stretcher. Photolithographic patterning of silicon wafers is carried out as described previously (Bhatia et al.,1998). Briefly, negative photoresist is coated on a silicon wafer, exposed to ultraviolet light (365 nm) through a transparency mask, and developed using high resolution printing of patterns with required width and separation between the parallel lines (10 u,m parallel lines and 10 UJTI spacing). Polydimethylsilioxane (PDMS) is prepared from a mixture of 2 liquid components (Sylgard 186 Kit, Dow Corning), poured onto developed wafer as a thin layer and after degassing, and cured. The resulting membranes are coated with the re uired ECMfor aligning Itie myocytes. Figure 13. Novel myocyte patterning assay for Membrane micropatteminn improved quantitative measurements of protein changes due to chronic cell stretching. A PDMS PDMS Mold mold is fabricated and used to make thin silicone membranes with the same alignment pattern. A collagen (or other substrate) layer is Silicone membrane applied to the surface, and myocytes applied to the membrane. The cells align with the membrane and take an elongated, rod-like Collagen coating phenotype, and stretch patterns (longitudinal and/or transverse) are then applied to these cell preparations. Myocytes align with pattern C3.1.h. Stretchers and Membranes. Silastic deformable membranes are prepared with a collagen type I and other substrate coatings similar to that used by Sadoshima et al. (Sadoshima et al.,1992). The cells (2 * 105 cells/cm2) are plated on patterned or unpatterned silicon membranes coated with collagen type-1 and equipped on equibiaxial (uniform) or non-equibiaxial (elliptical) stretcher devices as shown in Fig. 12B (Lee et al., 1996). The cells are serum-depleted for 24 hr followed by passive stretch for an additional 24 hr. Total RNA is extracted and subjected to RNA protection assay (RPA) using Direct RPA system (Ambion Inc.). Mouse BMP 216 Knowlton, Kirk U. coding sequences were amplified by RT-PCR and used as the template for a BMP riboprobe synthesis with mouse GAPDH as a control. Micropatterning allows aligned myocyte cultures to be subjected to non- equibiaxial stretch on the elliptical stretcher (Gopalan et al., 2003), which applies anisotropic strain to cells in a constant ratio (e.g. 2:1). If cells are patterned with their axis aligned with one axis of the elliptical stretcher, the ratio of principal strains is defined by the ratio of minor to major diameters of the ellipse. With these devices, the magnitude of strain is precisely controlled at any value and calibrated for each stretcher. C3.1.I. Cell Preparations for Histology. In order to evaluate the sarcomeric definition and organization in myocytes plated on gels of different rigidities, cells are fixed and stained for markers such as a-actinin at 48 hours and 7 days after initial plating. Cells are fixed by washing with cold (4¿C) PBS followed immediately by immersion in cold 0.75% glutaraldehyde for 20 minutes. Cells membranes are permeabilized with 0.1% Triton X100 (Sigma) for 5 minutes. Antibodies are diluted in 1% bovine serum albumin (BSA, Gemini) and added to the fixed cells for 60 minutes at room temperature. Secondary antibodies of tetra-rodamine isothiocyanate (TRITC)-conjugated goat anti-rabbit (Jackson Immunoresearch, West Grove, PA) are diluted in 1% BSA and added for 30 minutes. Cells are imaged with a confocal microscope at NCMIR or by phase-contrast and epifluorescence using our Nikon Eclipse TE300 inverted microscope and Photometries Cascade 51 2X CCD camera and acquired using Metamorph software (Universal Imaging Corporation). In addition, cell area and aspect ratio (long axis/short axis) are computed by segmenting cells with ImageJ software (NIH). C3.1.I. Ratiometric Imagine] of Intracellular Calcium Transients. Isolated cells are labeled with Fura-2 (Grynkiewicz et al., 1985). Fura-2 calcium indicator (Molecular Probes) is diluted to a concentration of 5 uM in a high-calcium Tyrodes buffer (130 mM NaCI, 5.4 mM KCI, 2 mM CaCI, 1 mM MgCI2, 0.3 mM Na2HPO4, 10mM HEPES, 5.5 mM Glucose at pH 7.4) and added at room temperature for 30 minutes. Cells are then washed 3 times with fresh Tyrodes buffer and placed in Tyrodes buffer for the duration of the experiment. Cells are imaged using a Nikon Eclipse TE3000 inverted microscope and stimulated to beat using the same procedure as in the traction force measurements. A phase-contrast image of the resting myocytes is taken in order to constrain the analysis of the image. Using a Lambda DG-4 high-speed filter changer (Sutler Instruments), cells are illuminated with alternating 380 nm and 340 nm light and the fluorescence at 510 nm is imaged using a Photometries Cascade 51 2X CCD camera and acquired using Metamorph software (Universal Imaging Corporation). The filters are switched and a new image is acquired every 15 ms for 4.5 seconds (300 images). In order to calibrate the Fura-2 response, an equal amount of high-calcium Tyrodes buffer containing 20 uM calcium ionophore-4-bromo-A23187 (Molecular Probes) is added to saturate the Fura bound to calcium and images are acquired at 340 nm and 380 nm excitation/ 510 nm emission. The media is removed and media containing 10% ethylenediaminetetraacetic acid (EDTA, Sigma) is added in order to chelate all calcium and result in a baseline Fura fluorescent signal. Again, images are acquired at a 340 nm and 380 nm excitation/ 510 nm emission. Images are analyzed using custom macros programmed in Metamorph or ImageJ (NIH) to split the image sequence of alternating images at 340 and 380 nm excitation into 2 stacks, compute the ratio of those stacks and compute the average value of that ratio for each image inside the cell outline for manually outlined cell areas. The concentration of free calcium in the cell is then computed using the following formula: -R] max / where kd is the dissociation constant (0.14 uM), Q is the ratio of the total average fluorescence inside the cell outline at 380 nm excitation, R is the ratio of fluorescence at 340 nm excitation to the fluorescence at 380 nm excitation in each image, Rmin is the same ratio upon addition of EDTA and f?ma* is the same ratio upon addition of the calcium ionophore. C3.1.k. FRET Imaging of Intracellular Signaling Events. Neonatal cardiac myocytes are transfected with a recombinant FRET reporter, such as the A-kinase activity reporter AKAR2 (Zhang et al., 2005; Zhang et al., 2001) 1 day after isolation using FuGeneS transfection reagent (Roche Diagnostics) and imaged 2 days after transaction (-5% transfection efficiency). For adult cells we will used viral-mediated gene transfer. AKAR2 is a recombinant protein composed of the yellow (YFP) and cyan (CFP) mutants of GFP, a protein kinase A substrate, and a phosphothreonine-binding domain. PKA-mediated phosphorylation of AKAR2 increases FRET 217 Knowlton, Kirk U. from CFP to YFP, allowing real-time fluorescence imaging of endogenous PKA activity by the yellow/cyan emission ratio. Fixed myocytes expressing reporter are labeled for a-actinin to confirm normal cell morphology. Cells are washed with Hanks buffered saline solution with 20 mM HEPES buffer (pH 7.2) 20 minutes prior to imaging and kept at 35 degrees C with a stage heater (Carel) during imaging. Transfected myocytes are imaged on a Nikon Eclipse TE300 inverted microscope equipped with a 60X Plan Apochromat objective, Photometries Cascade 512F CCD camera, and MetaFluor 6.2 software (Universal Imaging Corporation). Imaging is performed using a 430/25 nm excitation filter (for CFP) and simultaneously recording CFP (470/30 nm) and YFP emissions (535/30 nm) with a DualView emission splitter (Optical Insights; Chroma filters). Images are acquired with 1 s exposure every 5 s. To obtain emission ratio time-courses, YFP or CFP emission intensities for each image are averaged over a region of interest, background subtracted, and then yellow/cyan emission ratio is calculated, normalized by the ratio before reagent application. For emission ratio images, a similar approach is used on a pixel-by-pixel basis with a 5x5 pixel median filter using MetaMorph software (Universal Imaging Corporation). C3.2. Isolated Murine Trabeculae and Papillary Muscles C3.2.a. Muscle Isolation and Mounting. Male or female mice are anesthetized with Isoflurane. After cervical dislocation, the chest is opened and the heart is arrested by intracardiac injection of low calcium, high potassium cardioplegic solution. Then the heart is rapidly removed, cannulated, and perfused with modified Hepes buffered solution containing (mM): 137.2 NaCI, 15.0 KCI, 1.2 MgCI2, 2.8 mM Na acetate, 10 taurine, 1.0 CaCb, 10mM glucose, and 10mM Hepes in equilibrium with 100% O2. The right ventricle is opened and a right ventricular papillary muscle is dissected free by first cutting a small section of the valve, to which the muscle is attached by way of the cordea, and then cutting a small section of the septal wall, to which the muscle it attached at the base. Papillary muscles are chosen based on their geometry. Typically papillary muscles chosen for these studies are long, thin, and unbranched. Studies are not done on left ventricular (LV) papillary muscles because these muscles are thick and diffusion of oxygen and nutrients to the core of these muscles may be limited. Additionally, ANP and BMP induction may be effected by a decreased oxygen supply in LV papillary muscles and hence these specimens may not have a normal stretch induced response (Chen, 2005; Sabatine et al., 2004). The muscles are mounted in the cardiac tissue culture chamber, containing the same Hepes buffered solution used for the dissection. Muscles are mounted between a basket attached to a force transducer and actuator controlled micromanipulator, and a stationary titanium hook like extension. Oxygen is flow over the solution during the mounting procedure, to prevent hypoxia. All portions of the system, for experiments longer than 6 hours in time, are steam sterilized prior to each experiment. Additionally, all solutions are filter sterilized. The Hepes buffered solution is then exchanged for a modified M199 cell culture media containing (mM): 2.0 L- carnitine, 5.0 creatine, 5.0 taurine, 2.0 L-glutamine, 0.2% albumin, 100 IU/ ml penicillin, 0.1 mg/ml streptomyocin, 10mM Hepes, and 50 ug/ml of insulin in equilibrium with 5% CO2and 95% Oa.The temperature of the muscle chamber is maintained at 34¿C. The pH of the solution is maintained between 7.4 and 7.5. The modified Hepes buffered solution is exchanged for modified M199 media in steps, such that calcium concentration is slowly increased from 1.0 mM to 1.75 mM. Once the solution is replaced, the muscle is stimulated via the hook and a platinum electrode positioned within close proximity of the muscle's base. Muscles are left at slack length and stimulated at 0.2 Hz for one hour (Janssen et al., 1998). After the muscle has equilibrated, it is either stretched between 85-95% of Lmax, where Lmax is defined as the length of the muscle at which the muscle produces the greatest active force, or left at slack length for the duration of the experiment (2 hr, 5 hr, 12 hr). Muscle lengths are acquired with an LVDT in parallel with the micromanipulator. The surface of each muscle can be marked with titanium dioxide markers, such that local muscle deformations can be observed during the experimental procedure. Muscle dimensions are acquired by video capture, calibrated with a phantom of known dimensions placed at the focal plane of the specimen. C3.2.b. Mechanical Testing. Right ventricular papillary muscles are mounted between a stationary hook and an Isometric Harvard Apparatus force transducer (724490), which is capable of measuring force ranges between 0-0.5g and has an accuracy of ¿1% (<1 mg). The Harvard Apparatus force transducer is attached to a Newport 460P series, high precision, linear modular ball bearing stage, whose position in the x/y/z direction is controlled by a micromanipulator, which is connected to a Newport motorized actuator (CM-12CC), having a resolution less than 100 nm, an incremental motion less than 500 nm, and a speed range between 50-500 218 Knowlton, Kirk U. Urn/sec. In order to measure muscle lengths and distances by which the muscle is stretched with a precision on the order of 1 |xm, a LVDT (Omega LD310-10) having an accuracy of 400 nm and a linear range of 20 mm, is mounted in parallel with the actuator. Using this experimental setup, uniaxial muscle forces are recorded while pacing the muscle at 1 Hz and during continuous stretching of the muscle up to Lmax. Muscle lengths are simultaneously acquired and local deformations of the muscle are imaged by a CCD camera (COHU Inc.). Digital video recordings of muscle deformation are synchronized with acquired force data. Lagrangian uniaxial strain measurements (E=1/2(A2-1), where Ais the stretch ratio) are calculated with respect to the slack length of each muscle. Lagrangian stresses are calculated by dividing acquired force data by the initial cross sectional area of each muscle. C3.2.C. Real-Time PCR Procedures. At the end of each muscle experiment, the muscles are immediately submerged in RNA later solution, which functions to stabilize mRNA in intact tissue samples and contains RNase inactivating reagents (Quiagen). Right ventricular papillary muscles are completely isolated from their attachments, the valve and septal wall. Then the muscles are homogenized and RNA is extracted from these tissues, using Quiagen's RNeasy Micro RNA purification kit. Given that total RNA extraction from these small specimens is low (-10 ng/|jJ) and that DNase treatment has been shown to degrade and or destroy nucleic acid extracts, DNase treatment is not applied to RV papillary muscle specimens (Jemiolo and Trappe, 2004). Reverse transcription (RT) is accomplished using the Invitrogen Super Script III cDNA synthesis kit, which is optimized for low inputs of RNA. Finally, quantitative PCR is performed using an Applied Biosystems ABI 7700 real time thermal cycler, AB Taqman Universal Master Mix with UNG, with AB pre-made primers and taqman probes for ANP, BMP, and GAPDH. ANP and BNP gene expression is normally induced by increased mechanical loads in isolated cardiac myocytes as well as intact cardiac tissue (Ruwhof et al., 2000). GAPDH housekeeping gene expression does not change in these experiments. For each amount of RNA tested, duplicate Ct values are obtained and averaged. Then the AACt method is applied to the averaged Ct values obtained, in order to quantify the fold change of ANP or BNP relative to the unstretched control samples. Within each gene expression assay, appropriate controls are also included. No RT controls are performed for each amount of RNA, testing for any genomic DNA contamination. Also negative controls, where no cDNA is added to the reaction, are performed in order to test the purity of the enzyme master mix as well as the primer and probe mixture. Data are presented as mean ¿ 1SEM. Because mRNA concentrations are log-normally distributed and require nonparametric statistics, a two group t-test is performed on the ACt values or the raw output data obtained from the thermal cycler, between stretched and unstretched experimental groups. Significance is set at the P < 0.05 level, with appropriate corrections for multiple comparisons. C3.2.d. Antibody Staining and Confocal Microscopy. Papillary muscles that are not used for gene expression analysis are stretched to some known % stretch and fixed in 2% PFA (para-formaldehyde) diluted in 50% PBS (phosphate-buffered saline) and 50% modified Hepes buffered cardioplegic solution. After 10-20 minutes, the solution is replaced with 4% PFA in 100% PBS. Muscles are allowed to fix for an additional 10-20 minutes and then transferred into Tissue-Tek O.C.T. compound. They are frozen on dry ice and stored at -80¿C. Using a cryostat, tissue samples are then sectioned (10-15 u,m sections). For ANP protein analysis, tissue cross- sections are prepared, while for titin epitope, staining longitudinal sections are prepared. After a series of blocking and washing steps, tissue sections are then stained with appropriate antibodies (e.g. IgG antimouse ANP antibodies are used to look at ANP protein expression in stretched and unstretched specimens, IgG monoclonal anti-mouse a-actinin (Sigma) antibodies stain for sarcomeric z-disks, monoclonal antimouse IgM 9D10 antibodies stain for the PEVK region on titin (Hybridoma Bank), and polyclonal anti-goat IgG telethonin antibodies stain for Tcap (Santa Cruz Biotechnology), a protein adjoined to the N-terminal end of titin). Appropriate secondary antibodies conjugated to either Alexa 568 or Alexa 488 were also used. For titin epitope staining procedures, the tissues were double stained with either a-actinin and Tcap or a-actinin and 9D10. A Biorad confocal microscope is used to image the sections with filter sets appropriate for the excitation of Alexa- 568 and Alexa-488 simultaneously. Distances between titin epitopes and the z-disk were determined using FFT analysis as we reported earlier for skeletal muscle fibers (Shah et al., 2004). 219 Knowlton, Kirk U. C3.3. Isolated Perfused Whole Mouse Heart Preparations C3.3.a. Isolated Mouse Heart Preparations. Mice are administered an intraperitoneal injection of heparin(100 USP Units), anesthetized with isoflurane and sacrificed by cervical dislocation. The heart is rapidly excised, washed in cold cardioplegic solution, and the aorta cannulated on a 20-gauge stainless steel cannula. The remaining tissue surrounding the aorta is tied closed using 5-0 suture to seal all vasculature to the heart. The heart is mounted on a retrograde Langendorff perfusion apparatus (Radnoti). The coronaries are perfused with an oxygenated modified Krebs-Henseleit solution: 24.9 mM NaHCOs, 1.2 mM KHaPO^ 11.1 mM dextrose, 1.2 mM MgSO4, 4.7 mM KCI,118 mM NaCI, and 2.55mM CaCI2 (Grupp et al., 1993). The perfusate is bubbled with 95% O2-5% CO2 gas, maintained at 35-37¿C, and a flow rate of 1-2 ml/min is achieved by applying a constant pressure of 70 mmHg. The coronaries are cleared and the heart is allowed to beat freely and equilibrate for 15 minutes. Surface EGG electrodes are set in place and recordings are taken throughout the experiment. 95% 02 / 5% COj The heart is paced with a digital stimulator (DS8000, World Precision Instruments). It is paced epicardially from either Retrograde Perfuston the atria or ventricle with a bipolar platinum electrode at a Reservoir constant current two times threshold. For isovolumic Langendorff preparations, a fluid filled balloon is inserted into Antarograde the left ventricle through the mitral valve. The balloon, Perfusion Reservoir consisting of plastic wrap at the end of a 20-gauge blunt needle, is connected to a syringe infusion pump and inline with a fluid-filled pressure transducer. The balloon is inflated Compliance Chamber and deflated to an EDP of 30 mmHg to precondition the Moan Aortic P Measurement tissue. For the working heart preparation, the heart contracts Outflow Resistance against a resistive-compliant network consisting of a chamber f. Cardiac Output with an air bubble and a flexible tube with an adjustable clamp (Fig. 14).The heart is kept immersed in warm (37¿C) Collection Reservoir KHS throughout the experiment except when marker deformation is being videotaped. The pressure in the aortic Right Atrtal outflow line is monitored using a transducer (Millar Pacing Loads Instruments, Houston, TX). Video Camera Figure 14. Schematic of isolated ejecting heart apparatus in the VCR Core A lab. (From Karlon ef a/. (Karlon et al.,2000)) C3.3.b. Non-Homogeneous Epicardial Strain Analysis. Non-homogeneous epicardial strain distributions are mapped in the isolated mouse heart using high-resolution optical imaging of 50-100 small titanium dioxide markers arrayed on the epicardium with a short-bristled brush. In addition to LV strains, we have also used these methods in our lab to map regional distributions of septal strains in isolated working and non-working mouse hearts (Karlon et al.,2000) and to reconstruct right ventricular strains during systole (Lorenzen-Schmidt et al.,2005). A bipolar stimulating electrode will be used to pace the heart from the right atrium and a high- fidelity Millar pressure-volume catheter will be used to measure ventricular volume and pressure. A high- resolution CCD camera will be used to image the epicardium during isovolumic or ejecting contractions at full range of end-diastolic volumes. Markers will be tracked to sub-pixel accuracy throughout the cardiac cycle using the object tracking tools of the MetaMorph (Universal Imaging) software package. The markers are mapped to a bicubic Hermite finite element mesh and a least-squares fit of marker displacements is used to compute two-dimensional non-homogeneous strain fields using software developed in Dr. McCulloch's laboratory (Karlon et al.,2000; Mazhari et al., 1998). Briefly, the epicardial surface is modeled with 6-8 bi-cubic Hermite prolate spheroidal finite elements, and two-dimensional reference marker coordinates measured from the video images are projected on to the surface. The deformed coordinates are then projected on to the same material points in the models, which are then refitted by least squares resulting in a parametrically deformed mesh from which regional strains can be interpolated at each frame. This approach has also been used in conjunction with optical mapping of electrical activity in the isolated perfused rabbit heart (Sung et al.,2003). In addition to computing isovolumic strains (which are substantial), it is possible to obtain strains representative of ejection phase wall motions in the non-working heart, by using a reference state at an end-diastolic frame at mid-high ventricular volume and a deformed state at an end-systolic frames at lower ventricular volume. These 220 Knowlton, Kirk U. techniques can all be applied to the right ventricle as well as the left, which will be important for Project 2 (Chen). C3.3.C. Optical Mapping of Action Potential Propagation. A 0.8 ml bolus of the voltage-sensitive dye di-4- ANEPPS (25 uM) is injected into the perfusion line. The ventricular epicardium is illuminated by two 470 nm wavelength, 3.6-W 40-LED cluster bulbs (Ledtronics, Inc.). Fluorescence emission is passed through a >610 nm high-pass filter, focused with a fast 50 mm lens (1:0.95, Navitar) and recorded by a 12-bit charge-coupled- device (CCD) camera (CA-D1-0128T, Dalsa). Images of a 9x9 mm2 field of view are acquired at 950 frames per second and a spatial resolution of 64x64 pixels. The time series of fluorescent images provides temporal intensity signals at each pixel. Each signal is normalized with respect to baseline intensity (AF/F) and inverted to better represent a cardiac action potential. Spatial phase-shift and temporal median filtering (Sung, 2001) are implemented to reduce noise in the signal. Activation time is determined as the time of maximum slope (dF/dt) during the action potential upstroke. This provides an activation map over the ventricular epicardium showing the rate and path of action potential propagation. Spatial gradients of activation time are used to calculate local apparent conduction velocity vector at each pixel (Bayly et al., 1998). Additionally, the curvature of the activation wavefront is calculated as the spatial gradient of the normalized conduction velocity vector field. Time of repolarization is calculated by identifying the peak signal and determining when the action potential has recovered 20%, 50%, and 80% back to baseline. Action potential duration (APD) is then defined as the time difference between repolarization and activation. APD dispersion is determined as the standard deviation of APD over the surface of the heart. Attenuation of motion during optical mapping studies is necessary for the analysis of repolarization. Modified Krebs-Henseleit solution with 1 mM CaCb and 15 mM of the electromechanical uncoupler 2,3-butanedione monoxime (BDM) is perfused during data acquisition to eliminate motion artifact. However, it has been demonstrated that BDM prolongs action potential duration and slows conduction velocity in the isolated murine heart (Baker et al., 2004). We have recently developed a combination of ratiometry and a motion tracking algorithm to eliminate motion artifact without the use of chemical uncouplers. Since the emission spectrum of the potentiometric dye is wavelength shifted with altered membrane potential, upright and inverted optical action potentials can be recorded at green (560 nm) and red (620 nm) wavelengths respectively. A wavelength splitting optical setup allows us to record fluorescence emission at low and high wavelengths simultaneously using a single CCD camera, a technique originally applied to ratiometric microscopy (Kinosita et al., 1991). Image registration and ratio calculation of the two wavelengths amplifies the magnitude of the recorded optical action potential and corrects for differences in pixel intensity values due to heterogeneous dye loading or uneven excitation lighting. To correct for myocardial motion during contraction, a Lucas-Kanade optical flow algorithm is used to compute a vector field of frame-to-frame pixel displacements that is used to perform a non- rigid sub-pixel resolution transformation of deformed frames back to the reference configuration. Combined, these techniques effectively correct for tissue displacement and local motion artifact in the optical recordings. C3.3.d. Programmed Stimulation. Restitution kinetics and vulnerability to arrhythmia are assessed in the isolated mouse heart by programmed S1-S2 stimulation protocol. The isolated heart is paced from the left ventricular epicardium via a bipolar platinum electrode with a digital stimulator (World Precision Instruments, DS8000). The ventricle is paced at a basic cycle length of 200ms (S1-S1) for >20 beats followed by a premature stimulus (S2) (Baker et al., 2000; Lerner et al., 2000). The S1-S2 cycle length is decreased until the effective refractory period is reached, i.e. when the S2 stimulus no longer induces an action potential. During this protocol, optical mapping is used to analyze action potential morphology (amplitude and duration) and conduction velocity of the S2 induced action potential (Knollmann et al., 2006). The relationship between these measurements and the S1-S2 diastolic interval can be used to assess the restitution kinetics of the mouse heart. Premature action potentials during short S1-S2 cycle lengths can induce arrhythmias in susceptible media. The arrhythmia vulnerability of transgenic hearts can be assessed by the incidence of ventricular tachycardias during this protocol (Lerner et al., 2000). Additionally, burst pacing or a train of several high- frequency S2 stimuli can induce arrhythmias in the isolated heart (Fig. 15). C3.3.e. Electrical Space and Time Constants. The effective space and time constants directly aggregate myocardial electrical resistance and capacitance and are measured in the isolated mouse heart using similar methods to those of Poelzing et al. (Poelzing et al., 2005). A 125-micron diameter Teflon-coated platinum 221 Knowlton, Kirk U. unipolar electrode was used to pace the mid-lateral LV free wall. After steady-state pacing from the electrode (100 paced beats), a 1-mA cathodal stimulus was delivered for 250 ms, 35 ms after the final drive train pacing stimulus, during the refractory period of the previous action potential, resulting in a cathode-break stimulus upon release of the 250 ms pulse. Signals are normalized by baseline action potential amplitude so that the tissue stimulus response could be compared across space. The common mode signal (mean signal distal from the stimulus