A Review of Reagents for Fluorescence Microscopy of Cellular Compartments and Structures, Part II: Reagents for Non-Vesicular Organelles

A wide range of fluorescent dyes and reagents exist for labeling organelles in live and fixed cells. Choosing between them can lead to confusion, and optimization for many of them can be challenging. Presented here is a discussion on the commercially available reagents that have shown the most promise for each organelle of interest, including endoplasmic reticulum/nuclear membrane, Golgi apparatus, mitochondria, nucleoli, and nuclei, with an emphasis on localization of these structures for microscopy. Included is a featured reagent for each structure with a recommended protocol, troubleshooting guide, and example image. © 2023 Wiley Periodicals LLC. Basic Protocol 1: Endoplasmic reticulum and nuclear membrane labeling using ER-Tracker reagents Basic Protocol 2: Labeling Golgi apparatus using dye-labeled ceramides Basic Protocol 3: Labeling mitochondria using MitoTracker Red CMXRos Basic Protocol 4: Labeling nucleoli using SYTO RNASelect Green.

A Review of Reagents for Fluorescence Microscopy of Cellular Compartments and Structures, Part III: Reagents for Actin, Tubulin, Cellular Membranes, and Whole Cell and Cytoplasm

Non-antibody commercial fluorescent reagents for imaging of cytoskeletal structures have been limited primarily to labeling tubulin and actin, with the key factor in choice based mainly on whether cells are live or fixed and permeabilized. A wider range of options exists for cell membrane dyes, and the choice of reagent primarily depends on the preferred localization in the cell (i.e., all membranes or only the plasma membrane) and usage (i.e., whether the protocol involves fixation and permeabilization). For whole-cell or cytoplasmic imaging, the choice of reagent is determined mostly by the length of time that the cells need to be visualized (hours or days) and by fixation status. Presented here is a discussion on choosing commercially available reagents for labeling these cellular structures, with an emphasis on use for microscopic imaging, with a featured reagent, recommended protocol, troubleshooting guide, and example image for each structure. © 2023 Wiley Periodicals LLC. Basic Protocol 1: Actin labeling Basic Protocol 2: Wheat germ agglutinin conjugates for plasma membrane labeling Basic Protocol 3: Labeling tubulin microtubules with Tubulin Tracker Deep Red Basic Protocol 4: Labeling whole cells or cytoplasm with 5(6)-CFDA SE.

A Review of Reagents for Fluorescence Microscopy of Cellular Compartments and Structures, Part I: BacMam Labeling and Reagents for Vesicular Structures

Fluorescent labeling of vesicular structures in cultured cells, particularly for live cells, can be challenging for a number of reasons. The first challenge is to identify a reagent that will be specific enough where some structures have a number of potential reagents and others very few options. The emergence of BacMam constructs has provided more easy-to-use choices. Presented here is a discussion of BacMam constructs as well as a review of commercially available reagents for labeling vesicular structures in cells, including endosomes, peroxisomes, lysosomes, and autophagosomes, complete with a featured reagent, recommended protocol, troubleshooting guide, and example image for each structure. © 2023 Wiley Periodicals LLC. Basic Protocol 1: Delivering targeted fluorescent proteins using pre-made, high-titer BacMam constructs Alternate Protocol 1: Non-pseudo-typed BacMam viruses in standard cell types and pseudo-typed BacMam viruses in hard-to-transduce cell types Basic Protocol 2: Labeling endosomes: pHrodo™-10k-dextran Basic Protocol 3: Labeling peroxisomes: BacMam 2.0 CellLight™ Peroxisome-GFP Alternate Protocol 2: Labeling peroxisomes using antibodies Basic Protocol 4: Labeling autophagosomes: Transduction of cells with Premo™ Autophagy Sensor GFP-LC3B Alternate Protocol 3: Labeling autophagosomes using antibodies Basic Protocol 5: Labeling lysosomes: LysoTracker Red DND-99.

Superresolution microscopy with novel BODIPY-based fluorophores

Multicolor single-molecule localization microscopy (SMLM) expands our understanding of subcellular details and enables the study of biomolecular interactions through precise visualization of multiple molecules in a single sample with resolution of ~10–20 nm. Probe selection is vital to multicolor SMLM, as the fluorophores must not only exhibit minimal spectral crosstalk, but also be compatible with the same photochemical conditions that promote fluorophore photoswitching. While there are numerous commercially available photoswitchable fluorophores that are optimally excited in the standard Cy3 channel, they are restricted to short Stokes shifts (<30 nm), limiting the number of colors that can be resolved in a single sample. Furthermore, while imaging buffers have been thoroughly examined for commonly used fluorophore scaffolds including cyanine, rhodamine, and oxazine, optimal conditions have not been found for the BODIPY scaffold, precluding its routine use for multicolor SMLM. Herein, we screened common imaging buffer conditions including seven redox reagents with five additives, resulting in 35 overall imaging buffer conditions to identify compatible combinations for BODIPY-based fluorophores. We then demonstrated that novel, photoswitchable BODIPY-based fluorophores with varied length Stokes shifts provide additional color options for SMLM using a combination of BODIPY-based and commercially available photoswitchable fluorophores.

General Staining and Segmentation Procedures for High Content Imaging and Analysis

Automated quantitative fluorescence microscopy, also known as high content imaging (HCI), is a rapidly growing analytical approach in cell biology. Because automated image analysis relies heavily on robust demarcation of cells and subcellular regions, reliable methods for labeling cells is a critical component of the HCI workflow. Labeling of cells for image segmentation is typically performed with fluorescent probes that bind DNA for nuclear-based cell demarcation or with those which react with proteins for image analysis based on whole cell staining. These reagents, along with instrument and software settings, play an important role in the successful segmentation of cells in a population for automated and quantitative image analysis. In this chapter, we describe standard procedures for labeling and image segmentation in both live and fixed cell samples. The chapter will also provide troubleshooting guidelines for some of the common problems associated with these aspects of HCI.

Methodology for Quantitative Characterization of Fluorophore Photoswitching to Predict Superresolution Microscopy Image Quality

Single-molecule localization microscopy (SMLM) image quality and resolution strongly depend on the photoswitching properties of fluorophores used for sample labeling. Development of fluorophores with optimized photoswitching will considerably improve SMLM spatial and spectral resolution. Currently, evaluating fluorophore photoswitching requires protein-conjugation before assessment mandating specific fluorophore functionality, which is a major hurdle for systematic characterization. Herein, we validated polyvinyl alcohol (PVA) as a single-molecule environment to efficiently quantify the photoswitching properties of fluorophores and identified photoswitching properties predictive of quality SMLM images. We demonstrated that the same fluorophore photoswitching properties measured in PVA films and using antibody adsorption, a protein-conjugation environment analogous to labeled cells, were significantly correlated to microtubule width and continuity, surrogate measures of SMLM image quality. Defining PVA as a fluorophore photoswitching screening platform will facilitate SMLM fluorophore development and optimal image buffer assessment through facile and accurate photoswitching property characterization, which translates to SMLM fluorophore imaging performance.

A review of reagents for fluorescence microscopy of cellular compartments and structures, part I: BacMam labeling and reagents for vesicular structures

Fluorescent labeling of vesicular structures in cultured cells, particularly for live cells, can be challenging for a number of reasons. The first challenge is to identify a reagent that will be specific enough where some structures have a number of potential reagents and others very few options. The emergence of BacMam constructs has allowed more easy-to-use choices. Presented here is a discussion of BacMam constructs as well as a review of commercially-available reagents for labeling vesicular structures in cells, including endosomes, peroxisomes, lysosomes, and autophagosomes, complete with a featured reagent for each structure, recommended protocol, troubleshooting guide, and example image.

A review of reagents for fluorescence microscopy of cellular compartments and structures, Part III: reagents for actin, tubulin, cellular membranes, and whole cell and cytoplasm

Non-antibody commercial fluorescent reagents for imaging of cytoskeletal structures have been limited primarily to tubulin and actin, with the main factor in choice based mainly on whether cells are live or fixed and permeabilized. A wider range of options exist for cell membrane dyes, and the choice of reagent primarily depends on the preferred localization in the cell (i.e., all membranes or only the plasma membrane) and usage (i.e., whether the protocol involves fixation and permeabilization). For whole-cell or cytoplasmic imaging, the choice of reagent is determined mostly by the length of time that the cells need to be visualized (hours or days) and by fixation status. Presented here is a discussion on choosing commercially available reagents for these cellular structures, with an emphasis on use for microscopic imaging, with a featured reagent for each structure, a recommended protocol, troubleshooting guide, and example image.

A review of reagents for fluorescence microscopy of cellular compartments and structures, Part II: reagents for non-vesicular organelles

A wide range of fluorescent dyes and reagents exist for labeling organelles in live and fixed cells. Choosing between them can sometimes be confusing, and optimization for many of them can be challenging. Presented here is a discussion on the commercially-available reagents that have shown the most promise for each organelle of interest, including endoplasmic reticulum/nuclear membrane, Golgi apparatus, mitochondria, nucleoli, and nuclei, with an emphasis on localization of these structures for microscopy. Included is a featured reagent for each structure with a recommended protocol, troubleshooting guide, and example image.

Specificity, promiscuity and localization of ARF protein interactions with NCS-1 and phosphatidylinositol-4 kinase-III beta

ADP-ribosylation factor (ARF) proteins are involved in multiple intracellular vesicular transport pathways. Most studies have focused on the functions of ARF1 or ARF6 and little is known about the remaining ARF isoforms. Although the mammalian ARF proteins share a high degree of sequence identity, recent evidence has indicated that they may control distinct trafficking steps within cells. A unanswered issue is the degree of specificity of ARF family members for different interacting proteins. To investigate potential functional differences between the human ARF proteins, we have examined the localization of all human ARF isoforms and their interactions with two ARF1 binding proteins, neuronal calcium sensor-1 (NCS-1) and phosphatidylinositol-4 kinase-IIIbeta (PI4Kbeta). Use of a fluorescent protein fragment complementation method showed direct interactions between ARFs 1, 3, 5 and 6 with NCS-1 but at different intracellular locations in live HeLa cells. Photobleaching experiments indicated that complementation did not detect dynamic changes in protein interactions over short-time scales. A more specific interaction between ARFs 1/3 and PI4Kbeta was observed. Consistent with these latter findings ARF1 but not ARF5 or 6 enhanced the stimulatory effect of PI4Kbeta on regulated exocytosis, suggesting a specific role for class-I ARFs in the regulation of PI4Kbeta.

Calcium gradients and the Golgi

Changes in intracellular free calcium regulate many intracellular processes. With respect to the secretory pathway and the Golgi apparatus, changes in calcium concentration occurring either in the adjacent cytosol or within the lumen of the Golgi act to regulate Golgi function. Conversely, the Golgi sequesters calcium to shape cytosolic calcium signals as well as initiate them by releasing calcium via inositol-1,4,5-triphosphate (IP(3)) receptors, located on Golgi membranes. Local calcium transients juxtaposed to the Golgi (arising from release by the Golgi or other organelles) can activate calcium dependent signalling molecules located on or around the Golgi. This review focuses on the reciprocal relationship between the cell biology of the Golgi apparatus and intracellular calcium homeostasis.

Movement of calcium signals and calcium-binding proteins: firewalls, traps and tunnels

In the board game 'Snakes and Ladders', placed on the image of a pancreatic acinar cell, calcium ions have to move from release sites in the secretory region to the nucleus. There is another important contraflow - from calcium entry channels in the basal part of the cell to ER (endoplasmic reticulum) terminals in the secretory granule region. Both transport routes are perilous as the messenger can disappear in any place on the game board. It can be grabbed by calcium ATPases of the ER (masquerading as a snake but functioning like a ladder) and tunnelled through its low buffering environment, it can be lured into the whirlpools of mitochondria uniporters and forced to regulate the tricarboxylic acid cycle, and it can be permanently placed inside the matrix of secretory granules and released only outside the cell. The organelles could trade calcium (e.g. from the ER to mitochondria and vice versa) almost depriving this ion the light of the cytosol and noble company of cytosolic calcium buffers. Altogether it is a rich and colourful story.

Stable Golgi-Mitochondria Complexes and Formation of Golgi Ca2+ Gradients in Pancreatic Acinar Cells

We have determined the localization of the Golgi with respect to other organelles in living pancreatic acinar cells and the importance of this localization to the establishment of Ca(2+) gradients over the Golgi. Using confocal microscopy and the Golgi-specific fluorescent probe 6-((N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)amino)hexanoyl)sphingosine, we found Golgi structures localizing to the outer edge of the secretory granular region of individual acinar cells. We also assessed Golgi positioning in acinar cells located within intact pancreatic tissue using two-photon microscopy and found a similar localization. The mitochondria segregate the Golgi from lateral regions of the plasma membrane, the nucleus, and the basal part of the cytoplasm. The Golgi is therefore placed between the principal Ca(2+) release sites in the apical region of the cell and the important Ca(2+) sink formed by the peri-granular mitochondria. During acetylcholine-induced cytosolic Ca(2+) signals in the apical region, large Ca(2+) gradients form over the Golgi (decreasing from trans- to cis-Golgi). We further describe a novel, close interaction of the peri-granular mitochondria and the Golgi apparatus. The mitochondria and the Golgi structures form very close contacts, and these contacts remain stable over time. When the cell is forced to swell, the Golgi and mitochondria remain juxtaposed up to the point of cell lysis. The strategic position of the Golgi (closer to release sites than the bulk of the mitochondrial belt) makes this organelle receptive to local apical Ca(2+) transients. In addition the Golgi is ideally placed to be preferentially supplied by ATP from adjacent mitochondria.

NAADP mobilizes Ca2+ from a thapsigargin-sensitive store in the nuclear envelope by activating ryanodine receptors

Ca2+ release from the envelope of isolated pancreatic acinar nuclei could be activated by nicotinic acid adenine dinucleotide phosphate (NAADP) as well as by inositol 1,4,5-trisphosphate (IP3) and cyclic ADP-ribose (cADPR). Each of these agents reduced the Ca2+ concentration inside the nuclear envelope, and this was associated with a transient rise in the nucleoplasmic Ca2+ concentration. NAADP released Ca2+ from the same thapsigargin-sensitive pool as IP3. The NAADP action was specific because, for example, nicotineamide adenine dinucleotide phosphate was ineffective. The Ca2+ release was unaffected by procedures interfering with acidic organelles (bafilomycin, brefeldin, and nigericin). Ryanodine blocked the Ca2+-releasing effects of NAADP, cADPR, and caffeine, but not IP3. Ruthenium red also blocked the NAADP-elicited Ca2+ release. IP3 receptor blockade did not inhibit the Ca2+ release elicited by NAADP or cADPR. The nuclear envelope contains ryanodine and IP3 receptors that can be activated separately and independently; the ryanodine receptors by either NAADP or cADPR, and the IP3 receptors by IP3.

Generation and modulation of cytosolic Ca2+ signals in pancreatic acinar cells: techniques and mechanisms

Calcium is a ubiquitous signalling molecule, known to control a vast array of cellular processes. In order to retain stimulus fidelity, the cell encodes the increases in intracellular calcium in the form of oscillations that are regulated both temporally and spatially. Here, we review recent work, using the pancreatic acinar cell as a model system, on the mechanisms employed to generate and modulate cytosolic Ca2+ signals, and the technical advances that have made these studies possible.

Non-uniform distribution of mitochondria in pancreatic acinar cells

The distribution of mitochondria in pancreatic acinar cells was investigated using confocal fluorescence microscopy and transmission electron microscopy (EM). Acinar cells were studied either after enzymatic isolation or in small segments of undisassociated pancreatic tissue. Loading of isolated acinar cells with Mito Tracker Green or Red, a fluorescence mitochondrial probe, showed that mitochondria are predominantly situated in the perigranular, subplasmalemmal and perinuclear regions. Subsequent applications of EM fixatives induced a leak of the fluorescent indicator to the cytosol but did not change the distribution of mitochondria. EM was then performed on isolated acinar cells and on acinar cells of pancreatic tissue segments. The intracellular distribution of mitochondria was quantified by calculating the percentage of the cross-sectional area that was occupied by mitochondria. In isolated acinar cells the highest density of mitochondria was seen in the perigranular region, where mitochondria occupied 25.69+/-1.58% of the area, then the subplasmalemmal region with 12.61+/-0.77% and the perinuclear region with 9.07+/-0.97% ( n=26). Similar results were obtained from acinar cells of pancreatic tissue segments: the perigranular 22.9+/-1.95%, subplasmalemmal 12.45+/-0.78% and perinuclear regions 9.07+/-0.97% ( n=26). The outer mitochondrial membranes were frequently positioned close to membranes of the ER, which followed the outer contour of mitochondria. Mitochondria were never found in direct contact with the nuclear envelope: there were usually layers of ER between the mitochondrial and nuclear membranes. Subplasmalemmal mitochondria were found in a very close proximity to the plasma membrane with no ER layers between the mitochondrial and the corresponding plasma membranes. We conclude that in pancreatic acinar cells mitochondria are preferentially distributed to perigranular, subplasmalemmal and perinuclear regions and this distribution is not affected by isolation or fixation procedures.