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干货丨Western Blot条带灰度分析——Image J和Graphpad轻松搞定 - 哔哩哔哩

estern Blot条带灰度分析——Image J和Graphpad轻松搞定 - 哔哩哔哩干货丨Western Blot条带灰度分析——Image J和Graphpad轻松搞定艾美捷

关注专栏/干货丨Western Blot条带灰度分析——Image J和Graphpad轻松搞定干货丨Western Blot条带灰度分析——Image J和Graphpad轻松搞定

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关注WB是研究蛋白表达的一个经典方法。对于一些时间点或者是不同组织蛋白表达量的分析就涉及到量的变化。一些凝胶成像软件带有此分析工具，比如Quantity One，Bandscan，Gel-Pro Analyzer等成像系统专用软件。除了这些软件，还有一个比较简单的综合性质图像处理软件Image J可以很方便的进行灰度分析。而且Image J是开源性质的免费软件，可以在其官网直接下载使用。对Western Blot条带进行数值化有两种方法——灰度分析和光密度分析。灰度是计算机图像分析仪根据图像颜色深浅分为256个级别。灰度值越小物体颜色越深。光密度是指光线通过溶液或某一物质前的入射光强度IO与该光线通过溶液或物质后的透射光强度Ib比值的对数OD=log（IO/Ib）；图像分析中，入射光和透射光强度分别被切片上最亮区域的平均灰度值和待测目标平均灰度值取代，则图像分析系统的光密度值=log（切片标本最亮区域的平均灰度值（g0）/待测目标的平均灰度值（g））。光密度值越大，物体颜色越深，阳性物质相对含量越大。对于灰度值来说，切片标本制作时染色时间长短，以及测量时显微镜照明光源电压的大小对其影响很大。而光密度是一个比值。是通过计算一张切片标本最亮区域的平均灰度值与该切片标本中待测目标的平均灰度值的比值，再根据数学公式计算而来的，所以受切片标本染色时间长短及照明光源电压关系很小。然而有研究实践显示，灰度值与光密度值二者之间存在着线性关系，都可以用来进行WB定量分析。同样，Image J软件进行WB条带定量分析时也存在两种方法，此处只列举一种方法，具体步骤如下。1、下载安装，并打开Image J软件。2、导入WB条带图片。File→Open→ 弹出对话框→选择文件夹→找到WB条带。3、把图片转化成灰度图片：Image→Type→8-bit。这时WB图片将会随之改变。4、消除图片背景的影响。Process→Subtract Background。在弹出的对话框中，填上50基本就可以了，也可以稍高些，并勾上Light background，点击OK即可。此时图片背景会变白一些。5、设置定量参数。Analyze→Set Measurements。在弹出的对话框中勾选Area、Mean gray value、Min & max gray value、Integrated density。6、设置单位。Analyze→Set Scale。在弹出的对话框“Unit of length”后面填上“pixels”，其他的不用改动。7、把WB图片转换成亮带。Edit → Invert。8、选择菜单栏下的一系列不规则圆圈，根据情况选择一种。然后将圆圈手动拉倒第一条带，并尽量将条带都圈起来。9、点击菜单栏Analyze下拉出现的measurement，即可弹出你选定区域的灰度统计值。也可以点击快捷键（英文状态下的键盘m）。10、手动移动不规则圆圈至下一条条带，重复8、9步骤，直至所有条带都被测量。11、当测定完所有条带，选结果中的“Edit”的“Select All”，然后复制数据“IntDen”到Excel表即可进行分析。也可以直接在Results对话框中，选择File → Save as，直接导出excel表格。 12、在excel表格中将目的蛋白的灰度值除以内参蛋白的灰度值，进行归一化处理。13、下载安装，并打开Graphpad软件。14、在对话框左侧栏目“New table & graph”下选择“Column”。然后再右侧选择所要作的图形，以及是否显示误差棒等，具体如下图。15、在弹出的表格中，填上excel中的比值。16、然后点击菜单栏上Insert → New Graph of Existing Data... 。也可以在工具栏Sheet中的“New”条目下找到。（其实这步可以省略）17、在弹出的对话框“Create New Graph”中，填写新图名称，选择分析方法，图形，以及误差棒的表示方法，具体如下。（其实这步可以省略）18、此时，可以在工具栏下左侧的状态栏“Graphs”文件夹中找到所建新图“Date 1”。 19、此时可以对图形的外观进行修改，增加图片题目以及XY轴名称。直接在图上对应位置左键点击修改。20、也可以借助工具栏“Change”中的功能键对图片的柱形图颜色、宽度、高度、间隔等进行调整。21、然后点击工具栏“Export”将新建的柱形图导出。22、上面只是单因素的柱状图构建方法。如果是双因素，应该在一开始选择“Column”时选择“Grouped”。切记！！更多相关详情：5步教你跑出漂亮的大分子western-blot图诀窍：http://www.bio-review.com/western-blot-operating-steps/Western blot 条带分析：http://www.bio-review.com/western-blot-stripe/Western blot常见问题摘选：http://www.bio-review.com/western-blot-wb/本文禁止转载或摘编

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Western Blot条带灰度分析

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免费软件imageJ分析Western blot蛋白条带灰度值 - 知乎

免费软件imageJ分析Western blot蛋白条带灰度值 - 知乎首发于中山大学医学博士分享科研经验切换模式写文章登录/注册免费软件imageJ分析Western blot蛋白条带灰度值爱分享的Leo博士中山大学医学博士Western blot应该是医学生物领域基础文章必须要做的实验。关于western blot的实验方法以及注意事项，知乎链接传送门如下辛辛苦苦做出来了western blot条带，一定要让它在SCI文章中出现，载入史册。安装imageJ软件，官方软件下载网址链接传送门，是免费的哟安装完之后，咱们打开imageJ。界面虽小，功能强大。咱们就拿一个条带分析一下，一步一步操作，学不会您把我两个头给扭下来。1.打开图片：File----open2.转化为灰度图片：image——type——8-bit3、除背景：process——subtract background，选择504、设置参数：analyze——set measurements选择面积area平均密度mean gray value灰密度integrated density5.设置定量单位：analyze——set scale设置为pixel（像素）6、圈中条带：analyze——gels——select first lane7.分析圈中条带：analyze——gels——plot lanes8.得到结果选中直线工具，将开口的峰封闭；选中魔棒工具，点击封闭的峰即可显示该峰的面积，即为蛋白条带的灰度值9.均一化处理得到目标条带的灰度值之后，再使用相对应的内参灰度值进行均一化处理，即可得到最终结果。咱们看一下文献中的western blot结果是如何展示的。原始条带一张图，定量后扩成了三个图，数据一下子多了起来呢。编辑于 2022-04-26 14:10科学引文索引（SCI）医学药学赞同 11938 条评论分享喜欢收藏申请转载文章被以下专栏收录中山大学医学博士分享科研经验中山大学工作，中国科研人，

纯干货Western blot （WB）条带灰度统计与GraphPad作图 - 知乎

纯干货Western blot （WB）条带灰度统计与GraphPad作图 - 知乎切换模式写文章登录/注册纯干货Western blot （WB）条带灰度统计与GraphPad作图GBhouse攻击型人格请请请勿查看本人所有推文，谢谢！1 免疫印迹（western blot）灰度值（gray scale）计算实操1.1 image J 分析灰度值1.1.1 打开 image J；（图1 image J打开界面）1.1.2 打开需要分析的图片：“file- open- 找到图片”；（图2打开需要分析图片界面展示，此处展示的内参β-ACTIN）1.1.3 将图片转换为灰度图：“image- type- 8 bit”；（图3 western blot 图片转换为灰度图设置）1.1.4 将图片背景灰度统一：“process- Subtract Background”，然后将“rolling ball radius，参数设置为50”，复选框选中light background；此处的设置能够消除背景的影响；（图4 图片背景灰度统一设置）（图5 图片背景灰度统一设置参数调整）（图6 完成灰度背景统一设置后图片展示，可见背景颜色一致）1.1.5 翻转图片的黑白：“edit- invert”；（图7 图片黑白翻转设置）（图8 图片黑白翻转后展示）1.1.6 分析测量设置：“Analyze>Set Measurements，选择Area面积、平均灰度值Mean gray value、极大小灰度值 min& max gray value、积分灰度值Integrated dendity”；（图9 测量参数设置命令操作）（图10 参量参数复选框设置）1.1.7 将测量的单位设置为像素 pixels：“analyze- set scale”；（图11 测量单位定义为像素 pixels设置）（图12 设置为像素的相关设置，并定义到全部Global）2. 数据采集2.1 条带的选择和对应光密度的测量：图15红色箭头指向，分别表示举行，椭圆、和不规则图形，我一般比较喜欢用矩形选框；（图13 选择条带的框）2.2 测量印记的灰度值，使用矩形/椭圆/不规则选框选择需要测量的印记，然后使用快捷键（ctrl + M）或者菜单命令“analyze- measure”，尽量使用同一个框去测量所有灰度条带（不是必须）；（图14 选中需要测量的印记条带）2.3 获得对应条带数据；（图15 测量结果展示，红框标记部分IntDen即为本次测量的光密度数值）2.4 条带数据的保存；（图16 数据导出设置）（图17 导出数据命名设置）采用同样的办法，将目标蛋白的灰度值计算出来！3 GraphPad软件统计分析与作图3.1 首先使用excel对蛋白相对表达进行预处理，其中target protein 和 β-Actin的数据是通过image J测量获得的灰度值值，而relative expression数值来自于target protein灰度值/β-Actin灰度值；（图18 Excle预处理数据示意图）3.2 打开GraphPad，选择“column”，然后点击create即可；（图19 创建GraphPad 柱状图column操作）3.3 对新建的表格进行命名，左侧红框表示分析时间+分析内容；右侧红框上边是阴性组NC和处理组Treat，下边对应的数值是本次检测蛋白相对表达的灰度值（GraphPad 转置粘贴快捷键“ctrl + shift+ T”）；（图20 新建数据表格命名以及数据张贴）3.4 数据分析“analyze”;（图21 数据统计选框）3.5 统计方法的选择，在此，因只有两组数据，我们选择 t test，然后点击OK，即可；（图22 选择 t test操作）3.6 使用参数非配对t test，如图中红框所示；（图23 使用参数非配对t test）3.7 统计结果解释说明，P value表示统计差异（＜0.05即表示有统计学差异），P value summary这里的几个星标可以用于后期作图，一般用1-3颗星即可，significantly different是对P value的进一步说明；（图24 统计结果展示）3.8 作图，点击左侧Graphs对应的统计表格名称即可，后侧是提供的图片展示类型，一般差异较小的使用mean/median & error；误差线 error bar 可选择SD或者SEM;（图25 数据作图以及图片展示类型选择）（图26 图片根据默认获得的图片展示）3.9 图片美化3.9.1 首先将主要title和X title删除，并对X-Y周进行命名；（图27 删除多余title和X-Y轴命名后图片展示）3.9.2 将Y轴的数值从0开始展示：双击Y轴，然后将 automatically determine the range and interval前的√去除，然后range 复选框中的minimum设置为0；（图28 Y轴相关信息设置）（图29 完成Y轴设置后的图片展示）3.9.3 图片颜色调整，双击住状图的柱子，可根据图30进行调整，一般情况下，只需要调整柱子填充颜色即可；（图30 柱状图柱子信息调整窗口）（图31 住状图颜色调整完成展示）3.10 图片导出3.10.1 导出的选择：在菜单栏的export即可找到，如下图所示（图32 图片导出菜单命令）3.9.2 图片导出前，需要进行一定的设置，一般设置导出图片格式为tif，图片像素，图片命名，图片储存位置等，具体见下图解释，完成图片导出后，务必去看一下是否导出成功。（图33 图片导出参数设置）4 结束。（爱自己！！！做科研!!! 每文格言张小娴说的：“我没有很刻意的去想念你，因为我知道，遇到了就应该感恩，错过了就需要释怀。”如果有用，关注楼主GBhouse，点赞+讨论是博主持续更新的动力！！！）发布于 2023-10-24 15:10・IP 属地陕西免疫蛋白印迹赞同 544 条评论分享喜欢收藏申请

手把手教学——WB条带量化之ImageJ软件分析 - 知乎

手把手教学——WB条带量化之ImageJ软件分析 - 知乎切换模式写文章登录/注册手把手教学——WB条带量化之ImageJ软件分析实验老司机坚持分享实验知识一、背景介绍众所周知，当我们得到WB条带后，可以通过条带的粗细、长短、深浅来初步判断某一蛋白的表达情况，但出于实验的严谨性及客观性，我们更需要通过对WB条带进行量化而得到客观依据进一步佐证我们的肉眼判断结果。本文就ImageJ软件对WB条带的量化作一简单介绍，当然，实现WB条带量化的软件并非ImageJ一个，常用的还有Quantity One、Gel-Pro Analyzer 等，感兴趣的同学可以了解一下。二、操作流程方法一：1)打开ImageJ软件，点击File→Open→找到待分析条带的位置即可打开（或者直接将图片拖拽到该软件也可实现）； 2)点击Image→Type→8-bit，即可将图片转化为灰度图片（注：由于实验结果图片不便展示，所以此处仅以之前跑的不好的内参图做演示）； 3)点击矩形工具，框选所有条带（注：不包括Marker的条带哦）；4)点击Analyze→Gels→Select First Lane→Gels→Plot Lanes，即可得到峰图；5)点击直线工具，闭合每个峰的开口（注：一定要保证开口完全闭合，即直线拉至与横坐标相交；最边上的峰可画可不画，关系不大）； 6)选中魔棒工具，依次点击每个峰即可得到的Area值即为每个条带量化值。方法二：1)打开ImageJ软件，点击File→Open→找到待分析条带的位置即可打开（或者直接拖拽即可）；2)点击Image→Type→8-bit，即可将图片转化为灰度图片；3)去除背景：点击Process→Subtract Background→50.0pixels，勾选Light Background（注：50.0pixels为软件默认值，当然此处数值也可根据实验结果进行适当调整）；4)参数的设定：点击Analyze→Set measurement→勾选Area、Mean gray value、Min&max gray value、Integrated density；5)点击Analyze→Set scale→确保unit of length选项中的单位为pixel；6)点击Edit→Invert后选用椭圆形工具分割单个条带（注：也可采用其他形状工具，按个人习惯而定）； 7)点击Analyze→Measure，得到的Intden值即为WB条带量化值（注：每个条带依次测量，也可采用快捷键Ctrl+M实现）。小结：(1)简单来说方法一是一种较为快速粗糙的处理方式，而方法二则是在尽可能排除背景等因素的影响后更为精细的操作，两种方法均可实现对WB条带的量化，择其一对条带进行分析即可。(2)用于分析的条带最好摆正以便于分析，也就是说最好在显影的时候就一次性摆正条带，否则后期还需要用软件实现条带拉直等处理，比较麻烦。本文作者是"夏沫梦鸢"同学，在获得她的授权后实验老司机将本文发表于公众号。文稿：夏沫梦鸢校对：煲仔饭素材：Canva参考资料：https://images.app.goo.gl/vJ3ifRPRNFDVwz3i9发布于 2024-03-01 10:32・IP 属地安徽ImageJ生物实验分子生物学赞同 1添加评论分享喜欢收藏申请

关于Western blot的灰度分析？ - 知乎

关于Western blot的灰度分析？ - 知乎首页知乎知学堂发现等你来答切换模式登录/注册软件数据分析统计软件灰度蛋白质组学关于Western blot的灰度分析？1.对于Western blot目的蛋白条带的灰度分析有哪些好用的软件？ 2.如果只保存了16bit的图，哪些软件可以识别？ 3.在ImageJ中做灰…显示全部关注者31被浏览187,031关注问题写回答邀请回答好问题 1添加评论分享10 个回答默认排序酸菜科研等 2 个话题下的优秀答主关注QuantityOne，Bandscan，Gel-ProAnalyzer，TanonImage等成像系统专用软件都可以用作灰度分析。除了这些软件，用的最多的就是综合性质图像处理软件ImageJ。16bit的意思是说，能够显示出来的颜色的数量有16位数。8、24都是一样的。一般灰度分析的要求图片8bit就好，image J可以调整。image J灰度分析步骤：打开Image J软件，导入WB条带图片：File→Open→找到WB条带。把图片转化成灰度图片：Image→Type→8-bit。第一个矩形工具→选上所有条带→analyze→Gels→select first laneanalyze→Gels→plot lanes（也可以分开选取，如：框选第一个条带→analyze→Gels→select first lane→将第一个框拖移到余下条带→Gels→select next lane），出现山峰样图。选中直线工具，将开口波峰关闭选中魔棒工具（正数第七个），点击波峰，就得出所有area值。最后在excel表格中将目的蛋白的灰度值除以内参蛋白的灰度值，进行归一化处理。在GraphpadPrism软件中作柱状图即可。其实得出的area和Intden严格来说不是灰度值，而是面积，但用来量化WB条带是OK的。还有一种计算Intden的办法，是光密度值，也可以用来量化WB条带。知乎专属福利；助大家一臂之力、我为大家准备了一份基础实验protocol,细胞侵袭、细胞凋亡、细胞黏着、细胞周期等，不仅有细胞培养相关实验，还有包括不同研究水平实验技术Protocol，不同实验方法全流程，WB实验流程、注意事项、数据处理及写作、IHC实验流程、操作技巧、注意事项、图像分析等相关实验的详细步骤，全都是经过前辈们无数次验证过的，希望对大家的实验有帮助。点击下方链接可扫码添加酸菜老师助手，知乎私信不回问题：中国临床医生科研成长平台；小助手会免费赠大家一门医学SCI从入门到精通学习营，添加即送！编辑于 2023-06-21 10:57赞同 13214 条评论分享收藏喜欢收起科研根号三江苏赛尔普生物，流式实验、单细胞测序、蛋白芯片、免疫检测等关注蛋白灰度分析软件有：Image-Pro Plus，Quantity One，Image J等~~过来人把几个软件的使用总结分享给大家，可以少走弯路软件的安装包和软件教程，有需要的可以自取免费，亲测安装！56个科研人必装软件安装包（11月更新）实验数据/图片处理+文献管理+科研绘图+写作翻译+统计分析+办公常用链接：https://pan.baidu.com/s/1JkjM67dHoKsY3K5AkLGrTg?pwd=6868 提取码：6868 1、三个软件的优劣分析1）Image-Pro Plus 综合性能最好，操作简便结果可靠！因为它有强大的 IOD 算法，能准确反应蛋白灰度~~但需要靠魔棒或轨迹法选取目的蛋白区域，有一定的主观性，不过蛋白灰度测定本身就是半定量分析-----测量值本身就没有一定标准，不同软件测量值不同，即使相同软件重复测量数值也不一定相同，因此只要测定数值能反应目的蛋白变化趋势，测量值相对偏差不大即可~~~（2） Quantity One 有泳道-轨迹测定法，等高线/手绘选取目的区域测定法等方法。i.泳道-轨迹测定法：过程：先剔除一系列背景，再选择泳道（lane），然后分析条带（band）,最后使用高斯建模得出灰度分析值（Gauss Model trace）特点：感觉比较科学，重复性好，但十分繁琐，而且有时不能正确反应灰度（我亲测过）ii.等高线-手绘选取目的区域测定法：通过等高线或手绘选取目的蛋白区域，最简便，但重复性不太好。（3）Image J 最坑爹~~~，它用魔棒选取区域测定灰度，可是有时很难选中所需区域，而且方法繁琐，不推荐！！在此，只总结综合性能最好的 Image-Pro Plus（IPP）蛋白灰度分析方法~~~~2、mage-Pro Plus 分析蛋白灰度教程说明：目的蛋白的灰度=目的蛋白 IOD 值/ 相应内参 IOD 值1. 使用 Photoshop 剪切出目的区域（只含 western bolt 条带，如图），保存为TIFF 格式文件。2. Image-Pro Plus 打开文件，剔除背景：（1）放大镜放大目的区域（2） Measure---calibration---intensity---options----image,选择背景最大 value 值------ok-------ok----system----close(3) Measure---count/size---measure---select measurements，选择 IOD 为测量值，调整结果显示方式：Options-label style 改为 mesurement，选择 IOD 为显示值。（4）接下来有 2 种方法选择目的蛋白区域（即找出 AOI，area of interest, IPP 核心元件）：魔棒和手绘轨迹法。魔棒：首选方法，客观科学，适合于蛋白条带轮廓清楚且各蛋白条带不融合（若条带弥散，相互融合，则魔棒无法选择单一蛋白条带，这时就要用手绘通过肉眼选择目的蛋白区域）。手绘轨迹法：主观性较强，当魔棒无法选择单一条带时才使用。（5）count/size---edit-----convert AOI to object---得出 IOD 值（6）若测下一条带，则点 NEW AOI 按钮，再按以上方法选择 AOI(7) 可以 count/size---view---measurement data 显示或输出测量值图示过程如下：魔棒法手绘轨迹法得出测量 IOD 值：下图显示的即为目的蛋白IOD 值重复测量其他条带：相关介绍：推荐内容绝密！2023年国自然立项清单表可下载（生命科学部+医学科学部），附往年1000+份中标标书！如何查看国家自然科学基金的的摘要和下载结题报告？有哪些好用的zotero插件?有什么适合药学/生物方向科研小白的简单易学的科研绘图工具？有没有什么很好的科研作图软件？顶级图像分析软件，Image J、Fiji、Image pro plus，一次帮你搞定！ NoteExpress和EndNote相比哪一个更好用？（附安装包）2023年更新！EndNote X9永久激活版本（序列号激活），最稳定，可汉化！写的太全了，Endnote 插入文献以及调整参考文献格式，这是知乎上最好的教程！太牛了！Zlibrary回归，这里有最稳定的访问指南！western blot是为了什么？如何学习和记忆细胞信号通路？最值得推荐装！GraphPad Prism 9.3 ，功能更多！SPSS 27 安装激活，授权日期到期时间2037年编辑于 2023-12-11 16:39赞同 8添加评论分享收藏喜欢

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Introduction to Western Blotting

Image Analysis and Quantitation for Western Blotting

Overview

Sample Prep

Electrophoresis

Transfer

Immunodetection

Image Acquisition

Image Analysis

Image Analysis and Quantitation

Analyzing and quantifying your western blot images.

On This Page

Blot Quantitation and Background

Volume Box Tools

Lane and Band Tools

Normalization

Molecular Weight Determination

Analysis Tips

Resources

Blot Quantitation and Background

What Does Quantitation Mean?

Quantitating a western blot refers to the measurement of the signal emitted by your protein band(s) of interest. The signal intensity of the band is directly proportional to the concentration of your target protein. By analyzing the intensity of the signal, you may determine whether the expression of the target protein in one sample has increased or decreased relative to another sample or control. Note: western blots are semi-quantitative so they are ideal for showing relative protein expression rather than an absolute quantity.

Band Volume

A protein band is a feature that appears in a western blot image. A western blot image is made up of pixels, which contain information about how much signal was collected at each location in the image. A digital image of a blot can be thought of as data in three dimensions. Each pixel in a blot image has an x and y coordinate, in addition to an intensity value, which occupies the third dimension. Therefore, band quantity can be thought of as a volume, which is the total amount of signal for all the pixels within that band.

The pixels that comprise the part of a band have x and y coordinates, so outline an area. Each pixel also carries signal intensity information, which can be thought of as data in the third dimension.

Therefore, band quantity is often referred to as band "volume" by imaging software.

Blot Background

No western blot is perfect, and all blot images show some level of background. This background can come from non-specific binding during antibody incubation, membrane background in the case of fluorescent images, or background arising from the instrumentation itself.

For bright, isolated bands on smooth backgrounds, this process can be relatively straightforward. For situations where the background intensity is near that of the signal, or where the background is uneven, identifying which pixels belong to the band of interest, and identifying the relevant background to subtract can be challenging. Image analysis software typically offer tools to subtract background, depending on the method of band quantification used.

An intuitive way to think about background is to think about how we measure the size of mountains. Mountain height can be thought of in terms of their height above sea level, but if you want to climb a mountain, you are really interested in how tall the mountain is from its base.

With western blot bands, the "base" is the area where the band meets the membrane background. Membrane background is typically above the platten background because most experiments will have at least some intrinsic signal like autofluorescence. Calculating the band volume requires adding up all of the signal included in the band and subtracting signal contributed from the intrinsic background.

When the intensity of the background of the blot is near the intensity of the band of interest, or when the background is not uniform, isolating the signal away from background becomes more challenging.

Background on the blot or image can interfere with accurate quantitation of band signal and may prevent you from measuring the relative expression of your target protein. Image analysis software typically contain tools to help you reduce the impact of background on a band’s quantity and allow you to better quantify the target protein’s true signal. Note: If you have not selected your preferred background subtraction settings, then your software will automatically use default settings, which may or may not be optimal for your analysis.

Western Blot Image Analysis: Volume Box Tools

One of the most intuitive ways to quantify a band is to simply draw a box around it and quantify the signal within the box. Volume boxes should be drawn around the bands of interest in such a way that they include all of the intensity of the band with a minimal amount of surrounding background.

When using volume box tools, make sure to capture all of the band of interest without excessive background areas and without including adjacent bands.

Volume Box Tools: Background Subtraction

When using volume box analysis there are two common methods of subtracting background. The first is referred to as local background subtraction. In local background subtraction, the pixels immediately bordering the region of interest are averaged to establish the mean level of the background for the band of interest. Thus, each volume box has unique localized calculated level of background that is subtracted from it.

This method allows more customizable background subtraction, but it assumes that the background signal on the blot is additive with the signal from the band of interest. When comparing multiple bands that represent the same amount of protein, if one of the bands lies directly over a region with high background, then that specific band will result in a higher signal.

Another common method to subtract background is termed global background subtraction. In this method, you draw an additional region of interest and designate it as a background region that applies to the entire image. The mean value in this background volume box is then subtracted from all of the volume boxes, even if the background levels immediately surrounding the volume boxes vary.

Imaging software will typically report both the background subtracted volumes (called adjusted volumes) as well as the non-background subtracted volumes (simply called volumes).

Comparison of Global and Local Background Subtraction

A different background level is subtracted from each volume box. The pixels immediately bordering the outside of the volume box are averaged to establish the mean level of the background for that band of interest. Thus, each volume box has its own unique calculated level of background that is subtracted from it. This method is particularly useful when different bands are on areas of different background.

For this blot, global background subtraction is difficult because each band is sitting on a very different background environment and local subtraction would be a better choice.

Western Blot Image Analysis: Lane and Band Tools

The other common approach for quantitating western blots is commonly referred to as using lane and band tools. This approach utilizes a two-step procedure where each lane of a blot is identified in the image, and then individual bands are identified inside each lane.

The lane and band tools are particularly useful when you want to quantitate either the total protein signal in each lane or multiple bands in each lane. Lanes should be drawn around the entire region where bands are present and extend slightly outside the boundary to ensure that all protein signal in the lane is included. Once the lane frame is drawn, the lane profile can also be visualized. Additionally, lane and band tools use a different approach of background subtraction, one that makes it easy to apply a localized background subtraction to each band in a lane.

Overview of Lane and Band Workflow

The first step in using lane and band tools is to define the outlines of each lane on the blot.

Defining the lanes can be done automatically by the software, or by manually drawing lane frames and manually moving the lane outlines to follow the lanes on the blot.

If the lanes are not straight, or the gel has smiled or frowned, most software allows you to bend individual lane outlines to match the curve of the lane on the blot. Be careful to capture the entire lane, but avoid including neighboring lanes.

Next, the bands in each lane are identified. This can be as simple as clicking on the bands manually. Additionally, the software can detect bands automatically based on sensitivity settings set by the user. In either case, bands can be added, removed, or the boundaries of each band can be modified.

Most western blot image analysis software allows you to view a lane in profile, showing bands as peaks in a simulated electropherogram. The band boundaries can be precisely positioned in this view.

Lane and Band Tools: Background Subtraction

The final step in lane and band analysis is establishing the background to be subtracted. This is most easily visualized using the Lane Profile tool. In the Lane tool, background is typically determined using some variation of the rolling disc method.

In this method, it is helpful to imagine a disc of variable diameter (but always smaller than the largest blot feature) being “rolled” across the lane profile plot. The rolling disc will “fall up” the band peaks if the band peaks are wide enough and determine the background level. The disc size is adjustable, allowing more or less background to be removed. A small disc size fits further into band peaks and allows greater background subtraction. Conversely, large disc sizes will not fit very far into a band peak, reducing the amount of background subtracted.

This approach allows for a background subtraction that is based on the local background level and uses a single background subtraction setting that can be either applied to all lanes of a blot or to individual lanes.

The area in green shows the signal left after background subtraction. Left: A large rolling disc leaves most of the area in the bands untouched as it cannot “fall up” into each peak. Right: A small rolling disc travels up into the peaks and subtracts more signal. To determine if the subtracted background signal is sufficient or too much, an sideways image of a lane is positioned adjacent to the chromatogram.

Large Rolling Disc

Small Rolling Disc

Using a large rolling ball (center) leaves a lot of the background in the band volume (green area). A small rolling ball (right) does a good job of removing membrane background and leaving just the true signal from the bands.

Tip

For the most accurate quantitation, choose a disc size that provides consistent background subtraction between lanes.

Western Blot Normalization

Why Normalize?

At first glance, a western blot result like this would seem to indicate that the treatment lowered target protein expression.

But, if the lanes were not loaded equivalently, this conclusion cannot be made unless the difference in sample load is accounted for. With normalization, the true up or down regulation of the target protein can be inferred.

In western blotting, normalization refers to correcting data to reduce the effects of experimental error arising from variations in pipetting, sample concentration, or uniformity of protein transfer between gel and membrane. Normalization allows your western blot to be more quantitative and gives you confidence that differences in signal on a western blot reflect differences in protein expression between samples on the blot.

Normalizing western blot data relies on an internal loading control or normalization reference that is used to correct for these sources of experimental error.

There are two types of commonly used loading controls. The first control method is to use a protein that is present in all samples and its expression is not expected to be altered by the experimental treatment used in your experiment, often called a housekeeping protein. The second control method is to detect all proteins on the blot with a nonspecific agent to give the total protein signal.

In either case, any variation in the signal of the reference loading control indicates experimental variation rather than a difference in expression of the target protein. Regardless of which loading control is used, an effective normalization strategy requires two prerequisites to be met.

Accurate Normalization Requires:

The signal of the target protein and the loading control ought to be in their linear range, not saturated. Increases in protein levels in the samples must be detectable as differences in signal intensity on a western blot

The loading control must be unaffected by experimental perturbation. Misleading results will occur if an experimental treatment affects expression of the loading control

The Normalization Procedure

The correction underlying normalization is straightforward and some western blotting imaging software include tools to simplify the process. Whether you perform the process yourself, or rely on the software tools, the overall process is similar. A mathematical correction is used to compensate for sample-sample variability.

The first step is to quantify the background-subtracted protein signals for both the target protein as well as the loading control in each lane of the blot. Next, a reference lane is selected on the blot. This reference lane is usually the first sample lane or can be any other lane of your choice. Since the process of normalization is to essentially compensate for relative amounts of sample load, any lane can be selected as the reference. Then a normalization factor is determined by dividing the signal from the reference lane by the sample lanes to arrive at a normalization factor for each lane. This normalization factor is then applied to the protein of interest to the sample lanes by multiplying the signal from the protein of interest by the normalization factor.

Note: Some imaging software calculate the normalization factor by using the signal from the reference lane as the denominator. In this case, simply divide the signal from the protein of interest by the normalization factor.

Housekeeping Protein Controls

Housekeeping proteins are the most commonly used loading controls. These proteins are common to all samples in the experiment and are assumed to be unaffected by experimental conditions. Examples of commonly used housekeeping proteins include β-actin, GAPDH, and HPRT1 which are detected along with the protein of interest. Differences in the housekeeping protein signal indicate errors in loading the samples, non-uniform transfer, or other variabilities.

Normalization with Housekeeping Proteins

To perform normalization calculations, first quantify the signals from the protein of interest and the housekeeping protein for all lanes.

Then pick a lane to which you will use as the reference lane for the rest of the blot. Mathematically, any lane can be used as the reference, as the normalization factor will be applied to all other lanes as a ratio relative to the reference lane.

Then determine the normalization factor for each lane by deriving the ratio of the signals of the housekeeping protein in each lane, using the reference lane as the numerator. This ratio yields the difference in sample load between the reference and other lane.

Then apply the normalization factor to the protein of interest to each lane by multiplying the protein signal by the normalization factor for that lane. This gives you the corrected or normalized signal for your protein of interest in the other lanes on the blot. Now the relative expression of the proteins can be accurately compared between the lanes.

Limitations of Housekeeping Proteins

Accurate normalization using housekeeping proteins requires both of the following:

Signals are within their linear range

Their expression does not vary due to experimental conditions

Use of housekeeping proteins are vulnerable to both of these limitations, and care must be taken to ensure accurate results.

Varying Levels of Housekeeping Proteins

The expression level of housekeeping proteins may change due to:

Experimental conditions

Developmental changes

Post-transcriptional regulation

Cell differences

Tissue age and type

Using a housekeeping protein as a loading control requires that its expression remain constant across all samples of the experiment, and no change due to the experimental treatment. Published evidence shows several examples of differential expression of commonly used reference proteins (Ghosh R et al. 2014 and Eaton SL et al. 2013).

Therefore, before relying on a housekeeping protein as a loading control, validate that its expression is stable through the experiment and that it faithfully reflects the sample load.

Differences in five candidate housekeeping proteins and total protein staining between tumor and non-cancerous tissues in the validation sample set. Immunodetection measurements of housekeeping protein levels show poor linearity and do not accurately indicate cell lysate loading levels. Total protein normalization of HKPs yields consistent band intensities. Hu X et al. (2016). Common housekeeping proteins are upregulated in colorectal adenocarcinoma and hepatocellular carcinoma, making the total protein a better "housekeeper."

Linearity of tubulin, actin, and GAPDH in 0.5–4 μg of HeLa cell lysate. Note that the GAPDH signal is saturated and cannot be used for normalization.

Reliable normalization of target protein levels requires that the signals for both the target protein and the loading control are in their linear dynamic ranges. Unfortunately, housekeeping proteins are usually highly expressed, whereas target proteins are often expressed only in low abundance. Thus, to detect the target protein of interest, large amounts of cell lysate may need to be loaded, resulting in overloading of HKPs, yielding oversaturated reference bands, out of their linear range.

Total Protein as a Control

The use of total protein measurement for western blot loading controls (total protein normalization, TPN) is a method devised to minimize the inherent difficulties with using housekeeping proteins. Total protein levels can be determined by staining the membrane with total protein stains (e.g. SYPRO Ruby, Ponceau S, or Bio-Rad’s Stain-Free technology).

Because total protein stains are less sensitive than antibody-based immunodetection, they are far less likely to result in an oversaturated signal. As a result, they exhibit good linearity in the common loading range of 10–50 μg of cell lysate. Therefore, total protein normalization can be used successfully with most target proteins including those with low expression levels as both the target protein and the total protein signals will be within their respective linear ranges.

Since total protein normalization utilizes the entire protein load as the loading control signal, it is inherently less susceptible to expression variations of any one protein due to experimental conditions.

Stain-free imaging allows for total protein normalization.

Lane profiling of total protein signal helps remove background and refine band detection.

Advantages of Total Protein Normalization

Total protein normalization is less susceptible to experimental perturbation, so more faithfully reflects sample load

The linear dynamic range of most total protein stains better matches that of low expressing target proteins allowing for more accurate quantitation

Normalization with Total Protein Stains

In order to perform the normalization calculations, first quantify the signals from the protein of interest as well as the total protein signal in each lane. Software packages such as Image Lab Software contain tools for total protein normalization, simplifying this step.

Then determine the Normalization factor for each lane by deriving the ratio of the signals of the total protein in each lane, using the reference lane as the numerator. This ratio yields the difference in sample load between the reference and other lane.

Then apply the Normalization factor to the protein of interest to each lane by multiplying the protein signal by the Normalization factor for that lane. This gives you the corrected or normalized signal for your protein of interest in the other lanes on the blot. Now the relative expression of the proteins can be accurately compared between the lanes.

Learn More about Total Protein Normalization »

Molecular Weight Determination

Many blot analysis software packages allow you to estimate the molecular weight of your target protein. All that is required is that you include a molecular weight standard as one of the lanes on your blot and calculate a standard curve based on the known molecular weights of the proteins in the molecular weight standard.

A linear relationship exists between the logarithm of the molecular weight of a protein and relative front, Rf (the Rf is the ratio of the distance migrated by the protein to the distance migrated by the dye front). Plotting of the log of the known weights of the molecular weight standard as a function of Rf, describes a line with known slope and intercept and allows the calculation of the molecular weights of an unknown protein.

Calculating Molecular Weight

Molecular weight determination using SDS-PAGE gels takes advantage of the linear relationship between the logarithm of the molecular weight of a protein and relative front, Rf.

Measure the distance from the top of the resolving gel to each band of the standard and to the dye front.

Calculate the Rf (relative front) for each band in the standard:

Rf =

Distance to the band

Distance to the dye front

The resulting plot should give a line.

Calculate the Rf for the unknown band using the same formula

Plot the log (MW) as a function of Rf for the standards bands.

Generate the equation y = mx + b, and solve for y to determine the MW of the unknown protein.

Image analysis software typically contain tools that simplify the molecular weight calculations and only require you to identify what standard was loaded on the gel. You only then need to identify your bands of interest and set the location of the starting wells on the gel and location of the dye front.

Using Image Lab Software to Determine Molecular Weight

Most image analysis software contain tools for determining molecular weight. Here we use Image Lab Software as an example, but most software packages have similar tools.

The first step is to acquire the image of your blot. At least one standards lane should be included. For the most accurate results, it is best to load standards in both the first and last lanes.

Define the lanes on the gel, making sure to set the top of the lanes to be either the bottom of the wells or the top of the resolving gel. Set the bottom of the lane frame at the dye front.

Detect bands in the standards and sample lanes.

In the software, enter the type of standard used (or enter the molecular weight data for standards not listed in your software) and instruct the software which lanes are to be used as standard lanes.

The software will automatically plot the molecular weight as a function of Rf and interpolate the molecular weights of your proteins of interest.

In Image Lab, other regression methods such as point-to-point can also be used. This may provide a better fit for gels with non-uniform acrylamide or gels with some bands in the stacking gel.

Learn More about Image Lab Software »

Western Blot Image Analysis Tips

Optimize your western blots.

Faint bands or uneven background can make quantitative analysis challenging. The best practice to avoid these challenges is to optimize the upstream steps. Refer to previous sections of this guide to improve gel separation, clean up membrane background, and generate brighter target bands.

Validate your quantitation tools.

The best way to learn about your quantitation tools is to generate blots with known, titrated amounts of target protein. For example, run a blot where the lanes are loaded with 20 µg, 15 µg, and 10 µg of total protein. After analysis, the relative quantitation of the target bands should be 2, 1.5, and 1. Try different quantitation tools and settings until you can reliably reproduce the correct results.

Validate your loading control.

When using a housekeeping protein as a loading control for normalization, make sure its expression does not change due to the experimental conditions. If it does, then normalization will actually make your quantifying expression of your protein of interest less accurate. It is common to test multiple potential housekeeping proteins for a given experimental procedure.

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Total Protein Normalization

Compare changes in protein expression by establishing the baseline needed to correct against common errors such as inconsistent sample preparation, pipetting error, and uneven protein transfer.

Stain-Free Imaging Technology

Visualize proteins on gels and blots without staining and destaining steps, then use total protein normalization for quantitative western blots.

Fluorescent Western Blotting

Learn about advances in digital imaging and fluorescent western blotting that have resulted in overcoming common challenges in traditional western blotting.

Film vs. Digital Western Blot Imaging

Discover the advantages that digital imaging has over film.

Western Blot Doctor Troubleshooting Guide

Our self-help troubleshooting guide covers solutions to many common and not-so-common western blotting issues and helps your blots look their best.

Fundamentals of Western Blotting Course #5: Image Analysis

Documents

Protein Blotting Guide (PDF 7.2 MB)

Details on blotting technology, available products, and methods, plus tips, techniques, and troubleshooting.

Protein Electrophoresis Guide (PDF 8.5 MB)

A guide to polyacrylamide gel electrophoresis and protein detection, including theory, product selection, protocols, and more.

ChemiDoc MP System Brochure (PDF 1.5 MB)

Explore the features and benefits of the new high-end imaging system for the best fluorescence and chemiluminescence detection.

Stain-Free Approach to Western Blotting (PDF 864 KB)

Learn how to use stain-free technology for total protein normalization as an alternative to the standard blot normalization process.

Stain-Free Western Workflow Brochure (PDF 541 KB)

See how the Stain-Free Western Workflow compares to traditional western blotting workflows.

Determining the Appropriate Sample Load When Using a Stain-Free Western Workflow (67 KB)

Determine the correct sample load for the detection of target proteins before performing a western blot experiment.

Validating the Expression Consistency of a Housekeeping Protein (198 KB)

Discover how to validate the consistency of HKP expression before performing a western blot experiment.

Publications

A Defined Methodology for Reliable Quantification of Western Blot Data

Learn about the methodology to obtain reliable, quantitative data from chemiluminescent western blots using standardization procedures coupled with the updated reagents and detection methods.

The Design of a Quantitative Western Blot Experiment

Read a summary about a complete western blot workflow with a focus on sample preparation and data analysis for quantitative western blotting.

References

Ghosh R, Gilda JE, Gomes AV. The necessity of and strategies for improving confidence in the accuracy of western blots. Expert Rev Proteomics. 2014 Oct;11(5):549-60. PMCID: PMC4791038

Eaton SL et al. Total protein analysis as a reliable loading control for quantitative fluorescent Western blotting. PLoS One. 2013 Aug 30;8(8):e72457. PMCID: PMC3758299

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关于Western blot的灰度分析，这里有最全分析软件测评 - 哔哩哔哩

stern blot的灰度分析，这里有最全分析软件测评 - 哔哩哔哩关于Western blot的灰度分析，这里有最全分析软件测评科研根号三

关注专栏/关于Western blot的灰度分析，这里有最全分析软件测评关于Western blot的灰度分析，这里有最全分析软件测评

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关注蛋白灰度分析软件有：Image-Pro Plus，Quantity One，Image J等~~过来人把几个软件的使用总结分享给大家，可以少走弯路软件的安装包和软件教程，有需要的可以在公众号科研根号三回复关键词软件03领取！1、三个软件的优劣分析1）Image-Pro Plus 综合性能最好，操作简便结果可靠！因为它有强大的 IOD 算法，能准确反应蛋白灰度~~但需要靠魔棒或轨迹法选取目的蛋白区域，有一定的主观性，不过蛋白灰度测定本身就是半定量分析-----测量值本身就没有一定标准，不同软件测量值不同，即使相同软件重复测量数值也不一定相同，因此只要测定数值能反应目的蛋白变化趋势，测量值相对偏差不大即可~~~（2） Quantity One 有泳道-轨迹测定法，等高线/手绘选取目的区域测定法等方法。i.泳道-轨迹测定法：过程：先剔除一系列背景，再选择泳道（lane），然后分析条带（band）,最后使用高斯建模得出灰度分析值（Gauss Model trace）特点：感觉比较科学，重复性好，但十分繁琐，而且有时不能正确反应灰度（我亲测过）ii.等高线-手绘选取目的区域测定法：通过等高线或手绘选取目的蛋白区域，最简便，但重复性不太好。（3）Image J 最坑爹~~~，它用魔棒选取区域测定灰度，可是有时很难选中所需区域，而且方法繁琐，不推荐！！在此，只总结综合性能最好的 Image-Pro Plus（IPP）蛋白灰度分析方法~~~~2、mage-Pro Plus 分析蛋白灰度教程说明：目的蛋白的灰度=目的蛋白 IOD 值/ 相应内参 IOD 值1. 使用 Photoshop 剪切出目的区域（只含 western bolt 条带，如图），保存为TIFF 格式文件。2. Image-Pro Plus 打开文件，剔除背景：（1）放大镜放大目的区域（2） Measure---calibration---intensity---options----image,选择背景最大 value 值------ok-------ok----system----close(3) Measure---count/size---measure---select measurements，选择 IOD 为测量值，调整结果显示方式：Options-label style 改为 mesurement，选择 IOD 为显示值。（4）接下来有 2 种方法选择目的蛋白区域（即找出 AOI，area of interest, IPP 核心元件）：魔棒和手绘轨迹法。魔棒：首选方法，客观科学，适合于蛋白条带轮廓清楚且各蛋白条带不融合（若条带弥散，相互融合，则魔棒无法选择单一蛋白条带，这时就要用手绘通过肉眼选择目的蛋白区域）。手绘轨迹法：主观性较强，当魔棒无法选择单一条带时才使用。（5）count/size---edit-----convert AOI to object---得出 IOD 值（6）若测下一条带，则点 NEW AOI 按钮，再按以上方法选择 AOI(7) 可以 count/size---view---measurement data 显示或输出测量值图示过程如下：魔棒法得出测量 IOD 值：下图显示的即为目的蛋白IOD 值重复测量其他条带：本文禁止转载或摘编

灰度值分析

蛋白灰度分析

image j进行灰度分析

灰度分析怎么作图

灰度分析怎么分析

western blot灰度分析后如何作

灰度分析怎么选区域

wb灰度分析

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Overview

All iBright Imaging Systems feature on-board analysis software which is capable of many of the routine post-image capture analysis workflows, including molecular weight estimation, densitometry, and normalization. All analysis inputs are driven directly from the touchscreen interface.

Our standalone iBright Analysis Software is a free software for organizing and analyzing blot and gel images captured on iBright Imaging Systems. Our iBright Analysis Software was designed to build upon the on-board analysis workflows, with more functionality and with the added convenience of multiple software formats depending on preference and need.

The standard iBright Analysis Software is available in two formats: a desktop version (both PC and Mac OS compatible) and a cloud-based version accessible via web browser through our Connect platform. We now also offer a third version of iBright Analysis Software called iBright Analysis Software—Secure, designed from the desktop version of our software, and designed to support 21 CFR Part 11 compliance as part of our 21 CFR Part 11 compliance support package.

All versions of our iBright Analysis Software utilize four main workspaces: gallery, adjust, analyze, and annotate. Whether making a simple adjustment for a presentation, or to compare subtle differences between important samples, our software has you covered. We regularly update our software to add new features and to continuously improve the user experience.

Software workspace

Function

Features

Gallery

Explore and organize your images

View, store, sort, search, select, import, export, and create reports

Adjust

Fine-tune and refine your images for meetings and presentations

Rotate, straighten, flip, zoom, crop, invert, adjust contrast and gamma levels, apply false colors

Analyze

Gain a deeper understanding of the data

Auto-analysis (automatically detect lanes and bands), manual analysis (user-defined), quantitation (relative or absolute), and normalization using total lane protein and housekeeping proteins

Annotate

Label your image

Label analysis objects (Frames, lanes, bands & regions) through data table and/ or add text, boxes and arrows using annotation tools

On-board analysis software

Rather than having to move to a separate computer for post-image capture data analysis, all iBright Imaging Systems offer the convenience of on-board analysis software for routine analysis workflows. The on-board analysis software automatically identifies lanes and bands to streamline this often tedious step of analysis. Molecular weight marker overlay with molecular weight estimation, quantitation, densitometry and normalization analysis can be performed directly on the instrument. Up to four blots or gels can be analyzed simultaneously, greatly increasing throughput.

iBright Analysis Software

Manage your images with ease

The Gallery workspace can be sorted based on mode (application), date, or file name, so your data is readily retrievable and organized. Image preview is adjustable, so you can scan through closely related data without opening each image individually.

iBright Analysis Software—Desktop: Gallery workspace.

Analyze your data quickly

Image analysis, which can be complicated and cumbersome, can be simplified to only a few clicks in the Image Analysis workspace of iBright Analysis Software. Automatically analyze up to four mini blots or mini gels at a time from a single image for increased throughput. Powerful algorithms are designed to consistently and accurately select and identify lanes and band boundaries for molecular weight estimation, densitometry, or normalization. Adjustments can be made to band display sensitivity to fine-tune for the number of bands displayed for analysis. Images can also be analyzed manually if preferred.

iBright Analysis Software—Desktop: Image Analysis workspace. Blots analyzed using automatic analysis feature. Four separate blots are identified with four independent analysis frames that can be edited and analyzed separately.

iBright Analysis Software—Desktop: Adjust Image options are highlighted within the Image Analysis workspace, which can be used to straighten, crop, rotate, flip, or invert images according to need and preference.

iBright Analysis Software—Desktop: Image Analysis workspace. Use lane profile to adjust bands and rolling-ball background correction. Lane profile is the intensity profile plot of the selected lane. It is plotted with average pixel intensity of all pixels at that location or band if identified on the x-axis and pixel position on the y-axis. The chart also shows the non-specific background detected by the software in gray. By default, rolling-ball background subtraction is performed using an optimal rolling ball radius produced by the software for that image. This helps to more clearly view the band boundaries and more easily adjust the bands according to profile. Hovering over the profile graph provides values for band intensity and relative front.

Label for a more complete picture

The Annotate workspace allows the user to label the image. All annotations will appear in magenta in the viewport. Annotations and lane labels added using the annotation table will be included in the exported image and all annotations are included in the image report.

iBright Analysis Software—Desktop: Annotate workspace—lane and band labels. Lane labels were added through a data table and remaining labels were added using annotation tools.

Bring it all together with a complete report

After analysis, view your images alongside your data report. Data can be filtered to view only desired parameters. Reports can be exported to PDF format, or the raw data can be exported to Excel software for further analysis.

iBright Analysis Software—Desktop: Image report.

iBright Analysis Software—Desktop application

The iBright Analysis Software desktop application is available for anyone to download and is compatible with both PC and Mac operating systems. The desktop application allows you to work without the use of internet while still having access to all the capabilities of our cloud-based application.

Make sure your desktop software is up to date! Download the latest version in addition to instrument firmware updates and patches.

iBright Analysis Software—Cloud-based

Cloud connectivity helps increase productivity. Images captured on the iBright Imaging Systems can be exported directly to Connect, our cloud-based platform. Conveniently access via web wherever an internet connection is available. Furthermore, with Instrument Connect in Thermo Fisher Connect, you can determine instrument status, firmware edition, and usage history, providing an extra degree of control over the monitoring of your investment.

Each Connect account offers 1 TB of cloud storage at no charge. Link user accounts on an iBright Imaging System to a Connect account to directly upload data to a personal gallery on Connect.

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iBright Analysis Software—Secure

What is 21 CFR Part 11 Compliance?

The FDA released the Electronic Records and Signatures Rule, known as 21 CFR Part 11 in August 1997. This rule defines the requirements for use of electronic documents in place of paper documents. The law specifies the system elements, controls, and procedures that are necessary to ensure the reliability of electronically stored records.

21 CFR part 11 compliance is composed of both procedural and technical requirements. Procedural requirements are the standard operating procedures instituted by the end user, and technical requirements are the functional characteristics of the compliance management software used. Satisfying the technical requirements does not guarantee 21 CFR part 11 compliance alone. Compliance is the consequence of the end user’s work process and systems used.

For technical details on how the Invitrogen iBright SAE Software Solution for 21 CFR Part 11 Support helps enable workflows compliant to 21 CFR part 11, review the technical specifications document.

What is the iBright SAE Software Solution for 21 CFR Part 11 Support?

To implement the iBright SAE Software Solution for 21 CFR Part 11 support on iBright Imaging Systems, you need the following components installed, activated, and communicating with each other (Figure 1):

SAE Administrator Console Software: used with the iBright Imager and Analysis Application Profile to configure the security, audit and e-signature settings for iBright Imaging Systems

iBright SAE License: used to activate the SAE settings for the iBright instrument and the iBright Analysis Software—Secure desktop software

iBright Instrument—SAE Mode: firmware mode that connects the instrument to the SAE Administrator Console

iBright Analysis Software—Secure: desktop analysis software that connects with the SAE Administrator Console

Figure 1: iBright 21CFR11 Software Solution* comprised of (1) Security, Audit and e-signature (SAE) Software (SAE Administrator Console, SAE Server, and SAE Application Profile (i.e. iBright Imager and Analysis Software)), (2a) iBright Instrument – SAE Mode (onboard software), (3) iBright Analysis Software—Secure (desktop analysis software).

*iBright SAE Software License is required for activation of the 21 CFR Part 11 support package

SAE Administrator Console

The SAE Administrator Console is the tool used to configure security, audit, and e-signature settings to meet user-specific requirements for security, audit, and e-signature of a particular instrument and/or analysis software. The instrument and/or analysis software application profile (e.g. iBright Imager and Analysis Software application profile) must be installed in the SAE Administrator Console before the SAE settings can be used on the instrument and/or analysis software (Figure 2). Once the application profile is installed in the SAE Administrator Console, an SAE server is used to connect to the instrument and/or analysis software to the SAE Administrator Console. By default, the SAE server is installed on the same computer as the SAE Administrator Console, which should be a network-connected computer configured with a static IP address.

Figure 2: Interface view from the SAE Administrator Console

iBright SAE License

iBright SAE Software License for 21 CFR Part 11 is required to activate SAE Mode on the iBright Imaging System and to activate the iBright Analysis Software—Secure desktop analysis software.

iBright Imaging System SAE Mode

When SAE Mode is activated, the iBright instrument connects to the SAE Administrator Console via the SAE server. Security, Audit and e-signature settings, as defined in the SAE Administrator Console, are implemented on the iBright instrument (Figure 3). Features include restriction of unauthorized users to the system, password policies and expiration dates, defined user permissions and roles, tracking of all data changes through audit log, available audit reports, and on-board e-signature workflows. All iBright Imaging Systems can run in SAE Mode through purchase of the iBright SAE Software License.

Fig 3. Visuals from the iBright Imaging System interface showing how to access SAE mode.

iBright Analysis Software—Secure

iBright Analysis Software—Secure is a separate version of our iBright Analysis Software platform to support compliance with 21 CFR Part 11 FDA guidelines for Security, Auditing, and Electronic Signatures when connected to the SAE Administrator Console via the SAE server. The iBright Analysis Software—Secure desktop software can be utilized on computers running Windows or Mac operating systems, and can be configured to:

Control access to data using account credentials

Track user actions through audit logs

Detect data tampering

Sign electronic documents using electronic signatures

How does the iBright SAE Software Solution for 21 CFR Part 11 Support work?

Functional characteristics of the Invitrogen iBright SAE Software Solution for 21 CFR Part 11 Support

Function

Description

System security

Controls user access to an application by allowing creation of user account and defining user privileges through roles and defining password policies

Auditing

Tracks actions performed by users, changes to the SAE module settings, and creating audit reports

Electronic signature (e-signature)

Determines if users are required to fulfill signature requirements before performing specific functions

Additional design features for each function are detailed below:

System Security

1. Access is restricted to authorized personnel via user ID and password

Figure 4: Interface of the SAE Administrator Console showing new user account creation.

2. Passwords and user ID must expire after a user-defined number of days (typically 90 days).

3. Automatic logout after user-defined minutes of inactivity (Figure 5).

Figure 5: Interface of the SAE Administrator Console showing user name and password policy settings.

4. ID and password must be re-entered to sign or modify record.

5. Must have a method to record and automatically report ‘forgery’ attempts (event notification).

6. Access and manipulation of various software functions are controlled through user role definitions.

7. Control access to specific areas and options in software (Figure 6).

Figure 6: Interface of the SAE Administrator Console showing user role settings.

Auditing

Tracks actions performed by users and changes to the SAE settings. The SAE Administrator Console automatically tracks auditable actions silently (including, but not limited to, sign in/sign out, image acquisition, image exported, image deleted, file signed and report generation) (Figure 7).

Figure 7: Interface of SAE Administrator Console showing the audit history tab. (1) audit history available (action records, system configuration records, application object records), (2) application (SAE, iBright Instrument and Analysis Software, or other), (3) action, (4) date and user name (5) export in readable/printable format (PDF or .txt), (6) object ID.

With the auditing functionality, you can:

Select to audit specific user actions and specify the audit mode (silent, optional or required)

Generate reports for audited user actions and SAE module changes

Generate reports for software or instrument actions and events

Committing auditable actions

Auditable actions can be committed onboard the iBright Imaging Systems instrument or in the iBright Analysis Software—Secure software. In order to track changes, changes must be committed and saved. The committed changes made can be viewed under audit tab in the software application. Every commit creates a new version of metadata and is logged in the audit summary table in the audit tab. All committed actions are sent to SAE Administrator Console and logged under audit action records.

In the Commit workflow, select the Reason to commit from the selections displayed (or choose Other/Custom Reason). Enter comments as required and click on Commit (Figures 9 and 10).

Figure 9: iBright Analysis Software—Secure desktop software showing the reason to commit window and function (1) reason field (2) option to add a custom reason (3) comments.

Figure 10: iBright Imager SAE Mode auditable changes screens, displaying reason to commit window and function (1)reason field, touch “Blank reason” box to open audit reason screen (2) Select from options or “Other” to add a custom reason (3) enter comments if needed and touch Save, (4) confirmation screen of changes committed.

Electronic signature

Determines if users are required to fulfill signature requirements before performing specific functions (Figure 8).

Figure 8: Interface of the SAE Administrator Console e-signature settings. (1) customize the meanings of e-signatures (2) customize the number of e-signatures required by meaning and role (3) type of data signed for a specific selected meaning.

With the electronic signature functionality, you can:

Configure each e-signature event to require users with specific roles to sign

Create separate meanings with customized signature requirements for each meaning

Signing files and generating signed reports

When all the changes are done and committed, the file can be signed as per signature rules set in the SAE Administrator Console. Signature requirements (number of signatures and role required) for the selected meaning will be listed in the table (Figures 11a and 11b). Once all signatures are obtained, the status of the signature rule will be updated in the e-signature table for the corresponding version found in the audit tab from unsigned to signed. A signature log will be added in e-signature summary table.

In e-signature workflow, select the applicable e-signature meaning. Required signatures and roles will be listed. Select the applicable role and enter credentials. When all signatures are obtained, the file will be fully signed and locked.

Note: iBright Analysis Software—Secure allows partially signed files to be shared across user computers in partially signed state. When all signatures are obtained, the file will lock. iBright Instrument e-signature workflows requires all signatures to be obtained before exiting the e-signature workflow.

Panel A

Panel B

Figure 11: Signature workflow in iBright Analysis Software—Secure (panel A) and iBright Imager SAE Mode (panel B). Select Signature Meaning from available drop down. In iBright Analysis Software—Secure, each required role is listed, and signatures can be provided by clicking on “unsigned” status link for designated role. Partially signed files can be sent to additional users using iBright Analysis Software—Secure to complete all signature requirements. In iBright Imager SAE Mode, signature requirements for the designated Meaning are listed under the user icons. All signatures must be obtained at the same time onboard the iBright.

A report can be created directly from the iBright Imaging Systems instrument or from the iBright Analysis Software—Secure desktop software as either unsigned or signed. The file must be fully signed by all required roles/signatures prior to generating a signed report (Figure 12).

Figure 12. Export of image analysis report from iBright Analysis Software- Secure desktop software.

Documents

iBright Analysis Software User Guide

iBright Analysis Software Quick Start Guide

21 CFR Part 11

iBright SAE Software Solution for 21 CFR Part 11 Support—Welcome Letter

SAE Administrator Console User Guide

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