Correct localization and topology are crucial for a protein's cellular function. To determine topologies of membrane proteins, a new technique, called fluorescence protease protection (FPP) assay, has recently been established. The sole requirements for FPP are the expression of fluorescent-protein fusion proteins and the selective permeabilization of the plasma membrane, permitting a wide range of cell types and organelles to be investigated. Proteins topologies in organelles like endoplasmic reticulum, Golgi apparatus, mitochondria, peroxisomes, and autophagosomes have already been determined by FPP. Here, two different step-by-step protocols of the FPP assay are provided. First, we describe the FPP assay using fluorescence microscopy for single adherent cells, and second, we outline the FPP assay for high-throughput screening applications.

AMP-activated protein kinase (AMPK), a cellular metabolic sensor, is essential in energy regulation and metabolism. Hepatocyte polarization during liver development and regeneration parallels increased metabolism. The current study investigates the effects of AMPK and its upstream activator LKB1 on polarity and bile canalicular network formation and maintenance in collagen sandwich cultures of rat hepatocytes. Immunostaining for the apical protein ABCB1 and the tight junction marker occludin demonstrated that canalicular network formation is sequential and is associated with activation of AMPK and LKB1. AMPK and LKB1 activators accelerated canalicular network formation. Inhibition of AMPK or LKB1 by dominant-negative AMPK or kinase-dead LKB1 constructs blocked canalicular network formation. AICAR and 2-deoxyglucose, which activate AMPK, circumvented the inhibitory effect of kinase-dead LKB1 on canalicular formation, indicating that AMPK directly affects canalicular network formation. After the canalicular network was formed, inhibition of AMPK and LKB1 by dominant-negative AMPK or kinase-dead LKB1 constructs resulted in loss of canalicular network, indicating that AMPK and LKB1 also participate in network maintenance. In addition, activation of AMPK and LKB1 prevented low-Ca(2+)-mediated disruption of the canalicular network and tight junctions. These studies reveal that AMPK and its upstream kinase, LKB1, regulate canalicular network formation and maintenance.

Mitochondria are organelles that have been primarily known as the powerhouse of the cell. However, recent advances in the field have revealed that mitochondria are also involved in many other cellular activities like lipid modifications, redox balance, calcium balance, and even controlled cell death. These multifunctional organelles are motile and highly dynamic in shapes and forms; the dynamism is brought about by the mitochondria's ability to undergo fission and fusion with each other. Therefore, it is very important to be able to image mitochondrial shape changes to relate to the variety of cellular functions these organelles have to accomplish. The protocols described here will enable researchers to perform steady-state and time-lapse imaging of mitochondria in live cells by using confocal microscopy. High-resolution three-dimensional imaging of mitochondria will not only be helpful in understanding mitochondrial structure in detail but it also could be used to analyze their structural relationships with other organelles in the cell. FRAP (fluorescence recovery after photobleaching) studies can be performed to understand mitochondrial dynamics or dynamics of any mitochondrial molecule within the organelle. The microirradiation assay can be performed to study functional continuity between mitochondria. A protocol for measuring mitochondrial potential has also been included in this unit. In conclusion, the protocols described here will aid the understanding of mitochondrial structure-function relationship.

Rapidly emerging techniques of super-resolution single-molecule microscopy of living cells rely on the continued development of genetically encoded photoactivatable fluorescent proteins. On the basis of monomeric TagRFP, we have developed a photoactivatable TagRFP protein that is initially dark but becomes red fluorescent after violet light irradiation. Compared to other monomeric dark-to-red photoactivatable proteins including PAmCherry, PATagRFP has substantially higher molecular brightness, better pH stability, substantially less sensitivity to blue light, and better photostability in both ensemble and single-molecule modes. Spectroscopic analysis suggests that PATagRFP photoactivation is a two-step photochemical process involving sequential one-photon absorbance by two distinct chromophore forms. True monomeric behavior, absence of green fluorescence, and single-molecule performance in live cells make PATagRFP an excellent protein tag for two-color imaging techniques, including conventional diffraction-limited photoactivation microscopy, super-resolution photoactivated localization microscopy (PALM), and single particle tracking PALM (sptPALM) of living cells. Two-color sptPALM imaging was demonstrated using several PATagRFP tagged transmembrane proteins together with PAGFP-tagged clathrin light chain. Analysis of the resulting sptPALM images revealed that single-molecule transmembrane proteins, which are internalized into a cell via endocytosis, colocalize in space and time with plasma membrane domains enriched in clathrin light-chain molecules.

The endomembrane system of eukaryotic cells uses membrane-enclosed carriers to move diverse macromolecules among different membrane-bound compartments, a requirement for cells to secrete and take up molecules from their environment. Two recycling pathways-biosynthetic and endocytic, each with specific lipid components-make up this system, with the Golgi apparatus mediating transport between the two. Here, we integrate lipid-based mechanisms into the description of this system. A partitioning model of the Golgi apparatus is discussed as a working hypothesis to explain how membrane lipids and proteins that are segregated based on lateral lipid partitioning support the unique composition of the biosynthetic and endocytic recycling pathways in the face of constant trafficking of molecular constituents. We further discuss how computational modeling can allow for interpretation of experimental findings and provide mechanistic insight into these important cellular pathways.

Starvation-induced autophagosomes engulf cytosol and/or organelles and deliver them to lysosomes for degradation, thereby resupplying depleted nutrients. Despite advances in understanding the molecular basis of this process, the membrane origin of autophagosomes remains unclear. Here, we demonstrate that, in starved cells, the outer membrane of mitochondria participates in autophagosome biogenesis. The early autophagosomal marker, Atg5, transiently localizes to punctae on mitochondria, followed by the late autophagosomal marker, LC3. The tail-anchor of an outer mitochondrial membrane protein also labels autophagosomes and is sufficient to deliver another outer mitochondrial membrane protein, Fis1, to autophagosomes. The fluorescent lipid NBD-PS (converted to NBD-phosphotidylethanolamine in mitochondria) transfers from mitochondria to autophagosomes. Photobleaching reveals membranes of mitochondria and autophagosomes are transiently shared. Disruption of mitochondria/ER connections by mitofusin2 depletion dramatically impairs starvation-induced autophagy. Mitochondria thus play a central role in starvation-induced autophagy, contributing membrane to autophagosomes.

Recent developments in single-molecule localization techniques using photoactivatable fluorescent proteins have allowed the probing of single-molecule motion in a living cell with high specificity, millisecond time resolution, and nanometer spatial resolution. Analyzing the dynamics of individual molecules at high densities in this manner promises to provide new insights into the mechanisms of many biological processes, including protein heterogeneity in the plasma membrane, the dynamics of cytoskeletal flow, and clustering of receptor complexes in response to signaling cues. Here we describe the method of single-molecule tracking photoactivated localization microscopy (sptPALM) and discuss how its use can contribute to a quantitative understanding of fundamental cellular processes.