Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as addition/removal of synapses, occur in an experience-dependent manner, providing the structural foundation of neuronal plasticity. As postsynaptic components of the most excitatory synapses in the cortex, dendritic spines are considered to be a good proxy of synapses. Taking advantages of mouse genetics and fluorescent labeling techniques, individual neurons and their synaptic structures can be labeled in the intact brain. Here we introduce a transcranial imaging protocol using two-photon laser scanning microscopy to follow fluorescently labeled postsynaptic dendritic spines over time in vivo. This protocol utilizes a thinned-skull preparation, which keeps the skull intact and avoids inflammatory effects caused by exposure of the meninges and the cortex. Therefore, images can be acquired immediately after surgery is performed. The experimental procedure can be performed repetitively over various time intervals ranging from hours to years. The application of this preparation can also be expanded to investigate different cortical regions and layers, as well as other cell types, under physiological and pathological conditions.     

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of the olfactory epithelium continuously give rise to new sensory neurons that extend their axons into the olfactory bulb, where they face the challenge to integrate into existing circuitry. Because of this particular feature, the olfactory system represents a unique opportunity to monitor axonal wiring and guidance, and to investigate synapse formation. Here we describe a procedure for in vivo labeling of sensory neurons and subsequent visualization of axons in the olfactory system of larvae of the amphibian Xenopus laevis. To stain sensory neurons in the olfactory organ we adopt the electroporation technique. In vivo electroporation is an established technique for delivering fluorophore-coupled dextrans or other macromolecules into living cells. Stained sensory neurons and their axonal processes can then be monitored in the living animal either using confocal laser-scanning or multiphoton microscopy. By reducing the number of labeled cells to few or single cells per animal, single axons can be tracked into the olfactory bulb and their morphological changes can be monitored over weeks by conducting series of in vivo time lapse imaging experiments. While the described protocol exemplifies the labeling and monitoring of olfactory sensory neurons, it can also be adopted to other cell types within the olfactory and other systems.     

A Highlights from MBoC Selection: Molecular counting by photobleaching in protein complexes with many subunits: best practices and application to the cellulose synthesis complex

The constituents of large, multisubunit protein complexes dictate their functions in cells, but determining their precise molecular makeup in vivo is challenging. One example of such a complex is the cellulose synthesis complex (CSC), which in plants synthesizes cellulose, the most abundant biopolymer on Earth. In growing plant cells, CSCs exist in the plasma membrane as six-lobed rosettes that contain at least three different cellulose synthase (CESA) isoforms, but the number and stoichiometry of CESAs in each CSC are unknown. To begin to address this question, we performed quantitative photobleaching of GFP-tagged AtCESA3-containing particles in living Arabidopsis thaliana cells using variable-angle epifluorescence microscopy and developed a set of information-based step detection procedures to estimate the number of GFP molecules in each particle. The step detection algorithms account for changes in signal variance due to changing numbers of fluorophores, and the subsequent analysis avoids common problems associated with fitting multiple Gaussian functions to binned histogram data. The analysis indicates that at least 10 GFP-AtCESA3 molecules can exist in each particle. These procedures can be applied to photobleaching data for any protein complex with large numbers of fluorescently tagged subunits, providing a new analytical tool with which to probe complex composition and stoichiometry.     

Quantitative Analysis of Monocyte Subpopulations in Murine Atherosclerotic Plaques by Multiphoton Microscopy

The progressive accumulation of monocyte-derived cells in the atherosclerotic plaque is a hallmark of atherosclerosis. However, it is now appreciated that monocytes represent a heterogeneous circulating population of cells that differ in functionality. New approaches are needed to investigate the role of monocyte subpopulations in atherosclerosis since a detailed understanding of their differential mobilization, recruitment, survival and emigration during atherogenesis is of particular importance for development of successful therapeutic strategies. We present a novel methodology for the in vivo examination of monocyte subpopulations in mouse models of atherosclerosis. This approach combines cellular labeling by fluorescent beads with multiphoton microscopy to visualize and monitor monocyte subpopulations in living animals. First, we show that multiphoton microscopy is an accurate and timesaving technique to analyze monocyte subpopulation trafficking and localization in plaques in excised tissues. Next, we demonstrate that multiphoton microscopy can be used to monitor monocyte subpopulation trafficking in atherosclerotic plaques in living animals. This novel methodology should have broad applications and facilitate new insights into the pathogenesis of atherosclerosis and other inflammatory diseases.     

Fluorescent protein recruitment assay for demonstration and analysis of in vivo protein interactions in plant cells and its application to Tobacco mosaic virus movement protein

We describe a simple fluorescent protein‐based method to investigate interactions with a viral movement protein in living cells that relies on the in vivo re‐localization of proteins in the presence of their interaction partners. We apply this method in combination with fluorescence lifetime imaging microscopy (FLIM) to demonstrate that a domain of the Tobacco mosaic virus (TMV) movement protein (MP) previously predicted to mediate protein:protein interactions is dispensable for these contacts. We suggest that this method can be generalized for analysis of other protein interactions in planta.     

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisiæ, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.     

Induction of linear tracks of DNA double-strand breaks by alpha-particle irradiation of cells

Understanding how cells maintain genome integrity when challenged with DNA double-strand breaks (DSBs) is of major importance, particularly since the discovery of multiple links of DSBs with genome instability and cancer-predisposition disorders. Ionizing radiation is the agent of choice to produce DSBs in cells; however, targeting DSBs and monitoring changes in their position over time can be difficult. Here we describe a procedure for induction of easily recognizable linear arrays of DSBs in nuclei of adherent eukaryotic cells by exposing the cells to alpha particles from a small Americium source (Box 1). Each alpha particle traversing the cell nucleus induces a linear array of DSBs, typically 10-20 DSBs per 10 mum track length. Because alpha particles cannot penetrate cell-culture plastic or coverslips, it is necessary to irradiate cells through a Mylar membrane. We describe setup and irradiation procedures for two types of experiments: immunodetection of DSB response proteins in fixed cells grown in Mylar-bottom culture dishes (Option A) and detection of fluorescently labeled DSB-response proteins in living cells irradiated through a Mylar membrane placed on top of the cells (Option B). Using immunodetection, recruitment of repair proteins to individual DSB sites as early as 30 s after irradiation can be detected. Furthermore, combined with fluorescence live-cell microscopy of fluorescently tagged DSB-response proteins, this technique allows spatiotemporal analysis of the DSB repair response in living cells. Although the procedures might seem a bit intimidating, in our experience, once the source and the setup are ready, it is easy to obtain results. Because the live-cell procedure requires more hands-on experience, we recommend starting with the fixed-cell application.     

Viral nanoparticles as tools for intravital vascular imaging

A significant impediment to the widespread use of noninvasive in vivo vascular imaging techniques is the current lack of suitable intravital imaging probes. We describe here a new strategy to use viral nanoparticles as a platform for the multivalent display of fluorescent dyes to image tissues deep inside living organisms. The bioavailable cowpea mosaic virus (CPMV) can be fluorescently labeled to high densities with no measurable quenching, resulting in exceptionally bright particles with in vivo dispersion properties that allow high-resolution intravital imaging of vascular endothelium for periods of at least 72 h. We show that CPMV nanoparticles can be used to visualize the vasculature and blood flow in living mouse and chick embryos to a depth of up to 500 microm. Furthermore, we show that the intravital visualization of human fibrosarcoma-mediated tumor angiogenesis using fluorescent CPMV provides a means to identify arterial and venous vessels and to monitor the neovascularization of the tumor microenvironment.     

Dendra2 Photoswitching through the Mammary Imaging Window

In the last decade, intravital microscopy of breast tumors in mice and rats at single-cell resolution^1-4 has resulted in important insights into mechanisms of metastatic behavior such as migration, invasion and intravasation of tumor cells^{5, 6}, angiogenesis³ and immune cells response^7-9. We have recently reported a technique to image orthotopic mammary carcinomas over multiple intravital imaging sessions in living mice¹⁰. For this, we have developed a Mammary Imaging Window (MIW) and optimized imaging parameters for Dendra2¹¹ photoswitching and imaging in vivo. Here, we describe the protocol for the manufacturing of MIW, insertion of the MIW on top of a tumor and imaging of the Dendra2- labeled tumor cells using a custom built imaging box. This protocol can be used to image the metastatic behavior of tumor cells in distinct microenvironments in tumors and allows for long term imaging of blood vessels, tumor cells and host cells.Open in a separate windowClick here to view.^{(56M, flv)}     

Optimized Protocols for Mycobacterium leprae Strain Management: Frozen Stock Preservation and Maintenance in Athymic Nude Mice

Leprosy, caused by Mycobacterium leprae, is an important infectious disease that is still endemic in many countries around the world, including Brazil. There are currently no known methods for growing M. leprae in vitro, presenting a major obstacle in the study of this pathogen in the laboratory. Therefore, the maintenance and growth of M. leprae strains are preferably performed in athymic nude mice (NU-Foxn1^nu). The laboratory conditions for using mice are readily available, easy to perform, and allow standardization and development of protocols for achieving reproducible results. In the present report, we describe a simple protocol for purification of bacilli from nude mouse footpads using trypsin, which yields a suspension with minimum cell debris and with high bacterial viability index, as determined by fluorescent microscopy. A modification to the standard method for bacillary counting by Ziehl-Neelsen staining and light microscopy is also demonstrated. Additionally, we describe a protocol for freezing and thawing bacillary stocks as an alternative protocol for maintenance and storage of M. leprae strains.     

In vivo imaging of transplanted hematopoietic stem and progenitor cells in mouse calvarium bone marrow

In vivo imaging of transplanted hematopoietic stem and progenitor cells (HSPCs) was developed to investigate the relationship between HSPCs and components of their microenvironment in the bone marrow. In particular, it allows a direct observation of the behavior of hematopoietic cells during the first few days after transplantation, when the critical events in homing and early engraftment are occurring. By directly imaging these events in living animals, this method permits a detailed assessment of functions previously evaluated by crude assessments of cell counts (homing) or after prolonged periods (engraftment). This protocol offers a new means of investigating the role of cell-intrinsic and cell-extrinsic molecular regulators of hematopoiesis during the early stages of transplantation, and it is the first to allow the study of cell-cell interactions within the bone marrow in three dimensions and in real time. In this paper, we describe how to isolate, label and inject HSPCs, as well as how to perform calvarium intravital microscopy and analyze the resulting images. A typical experiment can be performed and analyzed in ～1 week.     

High-resolution Fiber-optic Microendoscopy for in situ Cellular Imaging

Many biological and clinical studies require the longitudinal study and analysis of morphology and function with cellular level resolution. Traditionally, multiple experiments are run in parallel, with individual samples removed from the study at sequential time points for evaluation by light microscopy. Several intravital techniques have been developed, with confocal, multiphoton, and second harmonic microscopy all demonstrating their ability to be used for imaging in situ ¹. With these systems, however, the required infrastructure is complex and expensive, involving scanning laser systems and complex light sources. Here we present a protocol for the design and assembly of a high-resolution microendoscope which can be built in a day using off-the-shelf components for under US$5,000. The platform offers flexibility in terms of image resolution, field-of-view, and operating wavelength, and we describe how these parameters can be easily modified to meet the specific needs of the end user.We and others have explored the use of the high-resolution microendoscope (HRME) in in vitro cell culture ^2-5, in excised ⁶ and living animal tissues ^2,5, and in human tissues in vivo ^2,7. Users have reported the use of several different fluorescent contrast agents, including proflavine ^2-4, benzoporphyrin-derivative monoacid ring A (BPD-MA) ⁵, and fluoroscein ^6,7, all of which have received full, or investigational approval from the FDA for use in human subjects. High-resolution microendoscopy, in the form described here, may appeal to a wide range of researchers working in the basic and clinical sciences. The technique offers an effective and economical approach which complements traditional benchtop microscopy, by enabling the user to perform high-resolution, longitudinal imaging in situ.     

Intravital multiphoton imaging of immune responses in the mouse ear skin

Multiphoton (MP) microscopy enables the direct in vivo visualization, with high spatial and temporal resolution, of fluorescently tagged immune cells, extracellular matrix and vasculature in tissues. This approach, therefore, represents a powerful alternative to traditional methods of assessing immune cell function in the skin, which are mainly based on flow cytometry and histology. Here we provide a step-by-step protocol describing experimental procedures for intravital MP imaging of the mouse ear skin, which can be easily adapted to address many specific skin-related biological questions. We demonstrate the use of this procedure by characterizing the response of neutrophils during cutaneous inflammation, which can be used to perform in-depth analysis of neutrophil behavior in the context of the skin microanatomy, including the epidermis, dermis and blood vessels. Such experiments are typically completed within 1 d, but as the procedures are minimally invasive, it is possible to perform longitudinal studies through repeated imaging.     

Monitoring Plasmid Replication in Live Mammalian Cells over Multiple Generations by Fluorescence Microscopy

Few naturally-occurring plasmids are maintained in mammalian cells. Among these are genomes of gamma-herpesviruses, including Epstein-Barr virus (EBV) and Kaposi''s Sarcoma-associated herpesvirus (KSHV), which cause multiple human malignancies ^1-3. These two genomes are replicated in a licensed manner, each using a single viral protein and cellular replication machinery, and are passed to daughter cells during cell division despite their lacking traditional centromeres ^4-8.Much work has been done to characterize the replications of these plasmid genomes using methods such as Southern blotting and fluorescence in situ hybridization (FISH). These methods are limited, though. Quantitative PCR and Southern blots provide information about the average number of plasmids per cell in a population of cells. FISH is a single-cell assay that reveals both the average number and the distribution of plasmids per cell in the population of cells but is static, allowing no information about the parent or progeny of the examined cell.Here, we describe a method for visualizing plasmids in live cells. This method is based on the binding of a fluorescently tagged lactose repressor protein to multiple sites in the plasmid of interest ⁹. The DNA of interest is engineered to include approximately 250 tandem repeats of the lactose operator (LacO) sequence. LacO is specifically bound by the lactose repressor protein (LacI), which can be fused to a fluorescent protein. The fusion protein can either be expressed from the engineered plasmid or introduced by a retroviral vector. In this way, the DNA molecules are fluorescently tagged and therefore become visible via fluorescence microscopy. The fusion protein is blocked from binding the plasmid DNA by culturing cells in the presence of IPTG until the plasmids are ready to be viewed.This system allows the plasmids to be monitored in living cells through several generations, revealing properties of their synthesis and partitioning to daughter cells. Ideal cells are adherent, easily transfected, and have large nuclei. This technique has been used to determine that 84% of EBV-derived plasmids are synthesized each generation and 88% of the newly synthesized plasmids partition faithfully to daughter cells in HeLa cells. Pairs of these EBV plasmids were seen to be tethered to or associated with sister chromatids after their synthesis in S-phase until they were seen to separate as the sister chromatids separated in Anaphase¹⁰. The method is currently being used to study replication of KSHV genomes in HeLa cells and SLK cells. HeLa cells are immortalized human epithelial cells, and SLK cells are immortalized human endothelial cells. Though SLK cells were originally derived from a KSHV lesion, neither the HeLa nor SLK cell line naturally harbors KSHV genomes¹¹. In addition to studying viral replication, this visualization technique can be used to investigate the effects of the addition, removal, or mutation of various DNA sequence elements on synthesis, localization, and partitioning of other recombinant plasmid DNAs.     

In vivo imaging of single axons in the mouse spinal cord

We provide a protocol that describes imaging of single fluorescently labeled axons in the spinal cord of living mice. This method takes advantage of transgenic mouse lines in which the thy1-promoter drives the expression of variants of the green fluorescent protein in a small percentage (less than 1%) of sensory neurons. As a consequence, single axons can be resolved in the surgically exposed dorsal column using wide-field epifluorescence microscopy. This approach allows direct observation of axonal degeneration and regeneration in mouse models of spinal cord pathology for several hours or repetitively over the course of several days.     

One-pot fluorescent labeling of xyloglucan oligosaccharides with sulforhodamine

Xyloglucan oligosaccharides fluorescently labeled with sulforhodamine have proved to be a valuable tool in the assessment of transglycosylating activity of plant xyloglucan endotransglucosylase/hydrolase (XTH; EC 2.4.1.207). Here we describe a simple and fast procedure for their preparation. Accordingly, the starting xyloglucan-derived oligosaccharides are in the first step converted to their corresponding 1-amino-1-deoxyalditols (glycamines) by incubation with ammonium acetate and NaCNBH(3) at 80 degrees C for 2-4 h, and in the second step, the glycamines are reacted with Lissamine rhodamine B sulfonyl chloride to obtain fluorescently labeled derivatives of the oligosaccharide glycamines. All operations are carried out in a single centrifuge tube and the products from the individual reaction steps are isolated on the basis of their differential solubility in organic solvents. Using the described protocol, the whole procedure can be accomplished in less than 24 h. The sulforhodamine-labeled xyloglucan oligosaccharides thus obtained proved suitable as substrates for a sensitive fluorescence assay of the transglycosylating activity of XTH.