High-speed imaging of FRET in live cells applied to investigate the role of PLCe in intracellular signal pathways

这项由伦敦帝国理工学院和癌症研究所联合发起的多学科联合项目旨在开发新技术，用于成像活细胞中蛋白质分子之间的相互作用，并应用于研究对癌症至关重要的细胞内信号通路。这些蛋白质相互作用将使用福斯特共振能量转移（FRET）的荧光技术成像。荧光成像需要用荧光分子（称为“荧光团”）“标记”感兴趣的蛋白质，荧光分子以特有的方式吸收和发射光。最近的一项突破是基因表达荧光蛋白的开发，这种荧光蛋白可用于标记活细胞中的特定蛋白质。传统的方法是用能被荧光团标签吸收的光子照射感兴趣的蛋白质来“激发”它们。然后它们会发出光（荧光）并放松到原来的状态。通过对这种荧光的强度成像，人们可以看到荧光团的分布，从而可以看到它们所附着的蛋白质。为了研究不同蛋白质之间的相互作用，可以用发出不同波长（即不同色光）的不同荧光团标记每种蛋白质。通过记录与每种蛋白质的分布相对应的不同颜色的图像，并将它们叠加在一起，人们可以看到不同蛋白质在同一位置出现的地方，即共定位。这种技术的问题在于，用于给活细胞成像的光学显微镜的空间分辨率受限于所讨论的光的波长（约400-700纳米），而蛋白质本身要小得多（约1-10纳米）。因此，即使两种蛋白质在荧光图像中出现在同一位置，它们在分子尺度上也可以是完全独立的。FRET提供了一种方法来确定当荧光团彼此在~ 10nm内/蛋白质相互作用的距离。它的工作原理是观察“激发能”从一个荧光团（称为“供体”）到另一个荧光团（称为“受体”）的转移，而不是只发生在很短的距离内。观察FRET最直接的方法是观察供体荧光强度降低或受体荧光强度增加的地方。不幸的是，由于背景噪声的影响，这种基于强度的成像往往不可靠。对FRET成像最可靠的方法是荧光寿命成像（FLIM）。一般来说，荧光寿命是通过用短脉冲光激发荧光团来测量的，并观察荧光信号在弛豫回基态时衰变所需的时间。使用非常快的相机技术，有可能成像荧光衰减在一个样品和获得荧光寿命值的每个像素在图像中。因为FRET提供了一个额外的方式，激发的荧光团失去他们的能量，可以确定FRET在哪里发生通过观察供体荧光寿命的减少。不幸的是，大多数FLIM技术相当慢，需要几分钟才能获得FLIM图像（荧光寿命值图），这使得很难使用FLIM- fret跟踪活细胞中的动态。该项目的目标是将帝国理工学院的新型高速FLIM和显微镜专业知识与ICR的生物专业知识结合起来，开发新的FLIM- fret成像系统。这将涉及设计具有合适的供体和受体荧光团标签的新分子，并将高速FLIM技术与新型显微镜配置相结合。这项提议的一个特别雄心勃勃的部分将是设计实验，在实验中我们可以进行多重FRET成像，即并行成像两种蛋白质-蛋白质相互作用，以研究细胞中的不同事件如何在信号通路中相关联。

This multidisciplinary joint proposal between Imperial College London and the Institute of Cancer Research is to develop new technology for imaging interactions between protein molecules in live cells, to be applied to study intracellular signal pathways that are important for cancer. These protein interactions will be imaged using the fluorescence-based technique of Forster Resonant Energy transfer (FRET). Fluorescence imaging entails 'labelling' proteins of interest with fluorescent molecules (called 'fluorophores') that absorb and emit light in a characteristic manner. A recent breakthrough has been the development of genetically expressed fluorescent proteins that can be used to tag specific proteins in living cells. Conventionally the proteins of interest would be 'excited' by irradiating them with photons that can absorbed by the fluorophore labels. These would then emit light (fluorescence) and relax back to their original state. By imaging the intensity of this fluorescence, one can visualise the distribution of the fluorophores / and therefore the proteins to which they are attached. To study interactions between different proteins, one can label each kind of protein with a different fluorophore emitting at a different wavelength (i.e. different colour light). By recording images of different colours / corresponding to the distributions of each kind of protein / and superimposing them, one can see where different proteins occur in the same place, i.e. co-localisation. The problem with this technique is that the spatial resolution of the optical microscopes that are used to image living cells is limited to ~ the wavelength of the light in question (about 400-700 nm) but the proteins themselves are much smaller (~ 1-10 nm). Therefore, even if two proteins appear to occur in the same place in the fluorescence image, they can be completely independent, on a molecular scale. FRET provides a way to determine when the fluorophores are within ~ 10 nm of each other / a distance at which the proteins would be interacting. It works by observing the transfer of 'excitation energy' from one fluorophore (called the 'donor' to another (called the 'acceptor') than only occurs over this very short distance. The most straightforward way to observe FRET is to see where the donor fluorescence intensity decreases or the acceptor fluorescence intensity increases. Unfortunately this kind of intensity-based imaging is often unreliable because of background noise. The most reliable way to image FRET is by fluorescence lifetime imaging (FLIM). In general fluorescence lifetime is measured by exciting fluorophores with a short pulse of light and observing how long it takes the fluorescence signal to decay away as they relax back to their ground state. Using very fast camera technology, it is possible to image fluorescence decays across a sample and obtain a value of fluorescence lifetime for each pixel in the image. Because FRET provides an additional way for excited fluorophores to lose their energy, one can determine where FRET is occurring by observing a reduction in the donor fluorescence lifetime. Unfortunately most FLIM technology is rather slow, taking several minutes to acquire a FLIM image (map of fluorescence lifetime values) and this makes it difficult to use FLIM-FRET to follow dynamics in live cells. The goal of this project is to combine novel high-speed FLIM and microscopy expertise at Imperial with the biological expertise at the ICR to develop new FLIM-FRET imaging systems. This will involve designing new molecules with appropriate donor and acceptor fluorophore labels and combining high-speed FLIM technology with novel microscope configurations. A particularly ambitious part of this proposal will be to design experiments in which we can 'multiplex FRET imaging, i.e. image two protein-protein interactions in parallel to study how different events in cells are associated in a signalling pathway.