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I've been doing a lot of live cell imaging lately mostly using hela cells expressing some EYFP based chimeric proteins. I'm building a video library for an art student here at the university who is going to create an exposition using the videos. I need a lot of content to make this sucessful and rather than requiring much scientific merit these videos need to be "busy" and look good. I'm looking for suggestions as to any dyes or fluorophores, drug treatments or conditions someone has worked with that will show significant movement/change of/in the cell, change in intracellular localization… doesn't matter it just needs to be a significant alteration for the cell.
I'm still really optimizing the system with the CO₂ etc… and the longest aquisition i have taken is about 8 hours - after that either the cells are dead or the focal drift becomes too much of a problem. I am hoping to see some division once I can get the system recording long enough. Until then was looking for some suggestions as to other possibilities.
The various dyes I have been using include: hoechst, dapi, mito-tracker, draq5, er-red, rox-red. also eyfp/tubulin, eyfy/mito, eyfp/golgi, draq5. These dyes aren't cheap and I'm only one i know doing this so I'm checking here.
MitoSOX looks pretty cool. I've never used it myself, but I'm sure you could see some cool mitochondrial dynamics going on. Its a bit pricey though, but all dyes are going to be.
Even at only 8 hours you should probably be able to see a few divisions, which would be cool with hoechst.
Cysteine (Cys), as one of the biothiols in organisms, is regarded as a significant participant in the physiological and pathological process. However, monitoring endogenous Cys in living organisms is still hindered due to the lack of effective fluorescent probe with excellent photophysical properties. Therefore, in this work, we attempted to develop excellent fluorescent probes for Cys detection with the aim of improving the photophysical properties of fluorophores. Firstly, through the rational structural modification of the dicyanoisophorone fluorophore by introducing different electron-withdrawing or electron-donating substituent groups, we found that three fluorophores with electron-withdrawing substituent group (-F, -Cl, -Br) exhibited remarkably decreased pKa value and complete deprotonation behavior in physiological pH, endowing the fluorophores with strong fluorescence emission in physiological condition and the wide pH adaptability. Subsequently, we developed three fluorescent probes (F-Cys, Cl-Cys, Br-Cys) for the Cys detection by introducing the acrylate group as the recognition unit into these three fluorophores, respectively. By comprehensively evaluating the sensing performance of three probes toward Cys, we screened out the best fluorescent probe Br-Cys. In solution, Br-Cys exhibited high sensitivity (detection limit: 86.9 nM), fast response (10 min), large fluorescence enhancement (214-fold), good selectivity, and wide pH adaptability. Furtherly, Br-Cys was successfully employed for the specific monitoring of Cys in cells and mice, providing a potential tool for understanding the physiological and pathological functions of Cys in living systems.
Fluorescent probe for in vivo detecting Cys based on the rational design of dicyanoisophorone through the strategy of lowering pKa.
Ehrlich’s use of synthetic dyes as a means of staining biological samples can be viewed as one of the foundation stones of modern scientific research. A century later, the use of fluorescence imaging as a technique to visualize specific regions of live cellular 1,2,3,4 or whole organisms 5,6 is often central to research programmes, with clinical applications such as fluorescence-guided surgery now emerging 7,9,10,11 .
The major shortcomings of fluorescence imaging using molecular fluorophores are interference from nonspecific background fluorescence outside the region of interest (ROI), insufficient photostability and cytotoxicity. Poor ROI selectivity necessitates a time delay to allow background fluorophore clearance and/or a washing procedure between fluorophore administration and image acquisition. This can limit imaging to fixed cells or static snapshots, without the possibility of continuous data acquisition throughout the experiment. An innovative approach to enhance target-to-background signal ratio is to exploit a mechanism of selective fluorescence quenching in the background areas, while establishing the emitting potential of the fluorophore only in the ROI 12,13 . Continuous recording of dynamic cellular events in real time may become feasible if the on/off fluorescence switching is reversible.
Developing a responsive fluorophore suitable for real-time live-cell imaging poses a series of challenges. Stringent criteria are required, such as near-perfect response selectivity, exceptional photostability and low dark and light toxicities. Obtaining selective fluorescence responses for intracellular analytes is not trivial, as analyte selectivity observed in a controlled homogeneous environment of a cuvette does not necessarily translate to far more complex in vitro or in vivo settings. Continuous live-cell imaging places very high demands on photostability of the fluorophore, as the same cell(s) are repeatedly imaged over time. Fluorophore dark toxicity must be low, so that cell viability is not compromised and normal cellular processes are unperturbed. To minimize light-induced toxicity, it is preferable to use low-energy wavelengths in the near-infrared (NIR) spectral region (λ=700–900 nm). For in vivo imaging, the use of NIR fluorophores is essential. This spectral region is required for effective light transmission through body tissue, as there are reduced levels of absorption and scattering at these longer wavelengths and less intrinsic autofluorescence. In addition, if on/off NIR fluorescence switching could be accomplished in vivo, then similar imaging advantages could be gained as for in-vitro cell imaging.
Currently, there is a small yet growing selection of NIR fluorophore classes but they often suffer from insufficient photostability and lack emission wavelengths above 700 nm 14 . Our recent research focus led to the development of BF2-chelated azadipyrromethene class 1 (Fig. 1) 15,16,17,18 . This class is relatively straightforward to synthesize, amenable to structural elaboration and exhibits excellent photophysical properties. For example, the derivative 1 (R=Ph) has an absorption/emission λmax at 696 and 727 nm in aqueous solutions, high fluorescence quantum yields (0.3–0.4) and excellent photostability 17 . Yet, in spite of recent progress, a significant need remains for new, more sophisticated intracellularly responsive molecular NIR fluorophores, which can be used to visualize dynamic cellular processes in real time with the potential for in vivo translation.
General structure of BF2-azadipyrromethenes 1. Design and synthesis of lysosomal responsive BF2-azadipyrromethene NIR fluorophore 2.
The goal of our current work was to develop an NIR fluorophore capable of a lysosomal-induced off-to-on fluorescence response, thereby permitting real-time imaging of cellular uptake, trafficking and efflux without perturbing function 19 . Endocytosis, the process through which cells internalize biomolecules, is common to all cells and represents a crucial area of research interest due to the numerous associated biological processes 20,21 . The participating organelles at each stage in the endocytosis pathway maintain a unique intravesicular/localized pH, to provide appropriate conditions for specific biochemical processes. Although the extracellular and cytosolic regions are at pH ∼ 7.2, the lysosomes are significantly more acidic. Along the endocytic pathway, the pH lowers from ∼ 6.3 in early endosomes through ∼ 5.5 in late endosomes, down to ∼ 4.5 in lysosomes (Fig. 2) 22 . As such, a difference of almost three orders of magnitude in proton concentration exists between the lysosome interior and the outside of a cell, which is sufficient to establish a selective trigger for fluorescence switching 23,24,25,26 . However, a major additional response selectivity challenge still remains, in that pH-responsive molecular fluorophores can also be responsive to micro-environmental polarity, which can compromise their use in cellular experiments (vide infra).
(a) Simplified endocytosis of a responsive NIR fluorophore. Numbers represent the approximate pH of the corresponding organelles. (b) Three observable stages of the path of the pH-responsive fluorophore in the cellular environment: uptake, trafficking and efflux.
Our novel lysosomal responsive probe design is illustrated in Fig. 1 in which functionalization of the fluorophore core (orange box) with an ortho-nitro phenolic group was chosen to impart the pH-responsive feature of the probe. It would be expected that the electron withdrawing o-nitro group would result in the ionized phenolate dominating at pH 7.2, resulting in fluorescence quenching due to a non-emissive intramolecular charge transfer excited state (Fig. 1, grey box). Following cellular uptake via endocytosis and compartmentalization into acidic organelles such as lysosomes, protonation would occur giving the neutral phenol species and the NIR emission signal would be established (Fig. 1, red box). This approach is a significant departure from other lysosomal stains, which rely on an amine protonation to form a positively charged ammonium salt to concentrate the fluorophore in the acidic compartments 19,22 . An important additional design feature includes a covalently linked polyethylene glycol (PEG) polymer to provide aqueous solubility and promote cellular uptake via endocytic pathways (Fig. 1, blue box) 27 .
Hela live cell confocal laser scanning - reccommendations for good fluorophore that will show good movement - Biology
An increasing number of investigations are using live-cell imaging techniques to provide critical insight into the fundamental nature of cellular and tissue function, especially due to the rapid advances that are currently being witnessed in fluorescent protein and synthetic fluorophore technology. Because of these advances, live-cell imaging has become a requisite analytical tool in most cell biology laboratories, as well as a routine methodology that is practiced in the wide ranging fields of neurobiology, developmental biology, pharmacology, and many other related biomedical research disciplines. Among the most significant technical challenges for performing successful live-cell imaging experiments is to maintain the cells in a healthy state and functioning normally on the microscope stage while being illuminated in the presence of synthetic fluorophores and/or fluorescent proteins.
Figure 1 - Live-Cell Imaging with Fluorescent Proteins and DIC
Tight control of the environment is one of the most critical factors in successful live-cell imaging experiments. In particular, the conditions under which cells are maintained on the microscope stage, although widely variable in many requirements depending upon the organism, often dictate the success or failure of an experiment. Aspects of the environment that are readily manipulated include the physical parameters of the chamber in which the cells are grown and imaged, the localized degree of temperature control, atmospheric conditions (gas mixture and humidity), nutritional supplements, growth medium buffering (pH), and osmolarity of the culture medium.
Illustrated in Figure 1 are a series of images captured from several unrelated cell lines, each labeled with a different combination of synthetic fluorophores and/or fluorescent proteins. The rabbit kidney epithelial cells (RK-13 line) in Figure 1(a) were transfected with a fusion plasmid of enhanced yellow fluorescent protein (EYFP) and a nuclear targeting signal peptide to localize a greenish-yellow label in the nucleus. The cells were subsequently treated with MitoTracker Red CMXRos to stain the mitochondria. In Figure 1(b), opossum kidney proximal tubule epithelial cells (OK line) were transfected with an EYFP-actin subcellular localization vector to label the filamentous actin cytoskeletal network. Mitochondria were targeted with a DsRed2 fusion vector in the Indian Muntjac cells presented in Figure 1(c), while an EGFP-peroxisomal chimeric plasmid highlights peroxisomes in the human cervical carcinoma (HeLa line) cells in Figure 1(d). The adherent culture of normal Syrian golden hamster kidney fibroblast cells (BHK-21 line) featured Figure 1(e) was transfected with a mixture of DsRed2 FP-endoplasmic reticulum and EGFP-nucleus subcellular localization vectors, thus localizing a green fluorescent protein tag to the nucleus and an orange-red probe to the endoplasmic reticulum. Finally, the human bone osteosarcoma cells (U2OS line) illustrated in Figure 1(f) was transfected with Cerulean fluorescent protein fused to a mitochondrial targeting sequence to label the mitochondria. For each of the images in Figure 1, a separate channel was recorded using differential interference contrast and overlaid on the fluorescence channel(s) to identify cell boundaries and other common structural features, such as the nucleus.
During the course of formulating plans for live-cell observations and long-term imaging experiments, a number of important factors are worthy of serious consideration. The specimen should be accurately labeled with the fluorescent protein or synthetic fluorophore(s) of interest in order to clearly visualize the target biological components. Perhaps even more important, the cell culture must be maintained in a condition that promotes growth and normal function in order to avoid potential artifacts in the interpretation of experimental results. In addition, the cells should be imaged with sufficient spatial and temporal resolution in a manner that does not induce phototoxicity or perturb localization of the fluorescent probes. Among the most important routine considerations for live-cell imaging that must be addressed (and discussed in detail below see Table 1) are temperature, oxygenation, humidity, osmolarity, pH (medium buffering), phototoxicity, the laboratory environment, microscope focus drift, fluorescence signal strength, bleed-through, and resolution.
Maintaining living cells in a healthy state on the microscope stage is undoubtedly the most critical aspect for any live-cell imaging investigation and generally requires a combination of mechanical ingenuity along with keen insight into the biology of the cell or tissue being studied. Although many laboratories are adept at growing cultured cells in temperature controlled carbon dioxide incubators, the task of maintaining cells on the microscope stage for long-term imaging experiments is far more demanding. The imaging chamber must keep the cells (or tissues) functioning normally for the duration of the experiment, while allowing unrestricted access by the microscope objective. This feat can become particularly difficult when high numerical aperture oil or water immersion objectives are being used. In many cases, the investigator must be able to introduce a reagent while imaging (to perturb a particular cellular process) without disturbing a time-lapse sequence by shifting focus or stage position. Other important factors are simplicity, reliability, and reasonable cost. The discussion that follows focuses primarily on mammalian cells, but the techniques for a variety of other organisms differ only slightly and are usually not as stringent. For example, cultures of yeast, insect, and plant cells do not have the strict temperature requirement, whereas the composition of media is not as demanding for bacterial cells.
Culture Media for Mammalian Cell Lines
Although the media for early attempts at cell and tissue culture consisted of mixtures containing embryo extracts, serum, protein hydrosylates and a host of other body fluids, propagation of established lines quickly required the transition into defined media based on biochemical requirements. Of these, the most popular are Eagle's Basal (EBM) and Minimal Essential (MEM) culture media, Dulbecco's modification of MEM (DMEM), Ham's media (F-10 and F-12), and two highly refined media formulations designed at the Roswell Park Memorial Institute (RPMI 1640 and RPMI 199). In addition, a medium designed for culturing cells in the absence of bicarbonate buffer (and carbon dioxide), Leibovitz L-15, has been widely employed. All of these culture media require the addition of serum (usually derived from fetal and newborn calves or horses) to a final volume fraction ranging between 5 and 20 percent. Serum-free media have also been developed for culturing highly specialized cells under strictly defined conditions that benefit applications in the biopharmaceutical industry. Many laboratories involved in culturing a wide variety of cell types often compromise on the rigorous requirements of an exact formulation by using a mixture of a complex medium, such as Ham's F-12, with a second medium (DMEM, for example) containing higher amino acid and vitamin concentrations.
The composition of cell culture media varies widely, but most recipes include amino acids, vitamins, inorganic salts (minerals), trace elements, nucleic acid constituents (bases and nucleosides), sugars, co-enzymes, lipids, tricarboxylic acid cycle intermediates, and a variety of other biochemicals. Simple media, such as MEM, contain only the essential amino acids, vitamins, and salts, whereas more complex formulations (RPMI and serum-free media) have hundreds of components. Cell culture media are usually designed for specific purposes, including routine growth of normal and immortalized (transformed) cell lines, primary culture initiation, virus propagation, pharmaceutical preparation, and defined growth conditions for genetic variants. Among the variables that are controlled in all tissue culture media formulations are pH, buffering capacity, oxygen concentration, osmolarity, viscosity, and surface tension. When cells are imaged in the microscope, even for short periods of time, these same medium conditions must be carefully reproduced in the live-cell imaging system.
Table 1 - Environmental Variables for Mammalian Cell Lines
|Temperature||28-37°C||Control with Specimen Chamber Heaters Use Inline Perfusion Heaters Objective Lens Heaters Environmental Control Boxes|
|Oxygenation||Variable||Perfuse or Change Media Regularly Use Large Chamber Volume|
|Humidity||97-100 Percent||Closed (Sealed) Chamber Humidified Environmental Chamber Auto-Fill System for Open Chambers|
|pH||7.0-7.7||Use HEPES Buffered Media Perfuse or Change Media Regularly No Phenol Red Indicator|
|Osmolarity||260-320 mosM||Avoid Evaporation Closed (Sealed) Chamber Humidified Environmental Chamber|
|Atmosphere||Air or 5-7 Percent Carbon Dioxide||Use HEPES Buffered Media for Air Closed (Sealed) Chamber Atmosphere Controlled Chamber|
|Media Buffer||Bicarbonate or Synthetic Biological Buffers||Beware of Phototoxicity Closed and Open Chambers Atmosphere Controlled Chamber|
A majority of the popular cell lines used in live-cell imaging experiments grow very well in a narrow range of pH between 7.2 and 7.4, although some normal fibroblasts perform better at slightly higher pH values (up to 7.7), while many transformed cell lines grow faster in more acidic media (down to pH 7.0). In cases where pH is critical for experimental accuracy, a plating efficiency assay should be performed at the target pH to ensure the selected cell line will perform satisfactorily. Most commercially available media formulations contain an indicator dye (usually Phenol red) for visual determination of the approximate pH value. In solution, Phenol red produces a bright red hue at pH 7.4, becomes orange at pH 7.0, and yellow at pH 6.5, a shift in color that is often seen when the medium becomes more acidic as cultures form confluent monolayers. At higher pH values, Phenol red is pink at pH 7.6 and purple at pH 7.8 and above. Many tissue culture laboratories find it useful to construct a set of pH standards using Phenol red in a balanced salt solution at varying pH values for comparison to culture media. Although the indicator dye is essential for routine cell culture, due to the high visible light absorption extinction coefficient, its use should be avoided in live-cell fluorescence imaging experiments in order to reduce the level of background noise and to prevent phototoxicity. Recognizing this fact, manufacturers provide most of the common media formulations in sterilized liquid or powder form without Phenol red.
Virtually all cell lines require a carbon dioxide and bicarbonate buffer system to regulate pH and must be cultured in an atmosphere containing a small percentage of carbon dioxide (usually 5 to 7 percent, depending upon the bicarbonate concentration) in specialized incubators to strictly control the concentration of dissolved gas. For live-cell imaging on the microscope, producing a suitable atmosphere with carbon dioxide can be difficult and usually requires culture chambers that are specifically designed for a regulated atmosphere. The introduction of synthetic biological buffers, such as TRIS and HEPES, has been of dubious value in eliminating the carbon dioxide requirement, as many cell lines will not tolerate a lack of carbon dioxide, especially at low cell concentrations. In general, a concentration of 10 to 20 millimolar HEPES buffer can control pH within the physiological range in the absence of a carbon dioxide atmosphere, but the culture medium should still be supplemented with sodium bicarbonate for optimum cell growth.
Attempting to grow cells on the microscope using HEPES alone usually results in dramatically reduced growth rates (especially for long-term experiments), and each cell line should be carefully scrutinized for its ability to grow and function in media without the carbon dioxide buffer system. Note that supplementing the bicarbonate buffer system in cell culture media with HEPES only reduces the rate of pH drift, and does not eliminate the progressive increase in alkalinity that occurs when the culture is exposed to the atmosphere. In addition, numerous reports have surfaced of HEPES toxicity during live-cell imaging experiments, presumably due to increased free radical formation by the synthetic buffer under the illumination conditions required to visualize fluorescent probes. A specialized formulation, Leibovitz L-15 medium, is designed to eliminate carbon dioxide through the use of sodium pyruvate and buffering with high amino acid concentrations. Including sodium pyruvate in the culture medium allows cells to increase their endogenous production of carbon dioxide, theoretically rendering them independent of the gas (as well as bicarbonate). However, many cell lines do not adapt well to L-15 medium and should be thoroughly examined for several passages before configuring live-cell imaging experiments based on this formulation.
Cell lines can vary widely in their oxygen requirement, although normal atmospheric oxygen tension levels will meet the needs of most cultures. Mammalian cells usually require oxygen for respiration in vivo, but can often successfully substitute glycolysis (an anaerobic process) when grown in culture vessels as primary lines or after immortalization. The depth of the culture medium above the cells can influence the rate of oxygen diffusion to adherent cells growing on a glass or plastic surface and should be kept below 5 millimeters. In most cases for live-cell imaging, strict oxygen regulation is not necessary and, conversely, depletion of oxygen is often used as a strategy to reduce the photodamage during fluorescence illumination that can occur through reactions with oxygen free radicals. However, it should be noted that lowering oxygen tension can be just as deleterious to cells if they begin to suffer hypoxic stress. The most common method of reducing oxygen levels involves the application of a commercial oxygen depletion system, such as Oxyrase. Alternative techniques to limit free radical damage due to molecular oxygen include supplementing the medium with scavengers such as ascorbate (ascorbic acid vitamin C) or Trolox (a derivative of vitamin E), but the strategy of reducing illumination intensity coupled with highly sensitive camera systems should also be considered.
Most of the popular cell lines have a fairly wide tolerance for osmotic pressure and will grow well at osmolarities between 260 and 320 milliosmolar. In cases where cells are routinely grown in Petri dishes or open-plate cultures, hypotonic medium can be substituted to compensate for evaporation. It is important to monitor osmolarity of the culture medium when altering the constitution by the addition of organic buffers or plasmid selection drugs, such as HEPES and G-418, respectively. The concentration of ions and organic nutrients in live-cell imaging will initially be set by the medium chosen for the experiment, but the small volumes of media accommodated by most imaging chambers are subject to changes in osmolarity due to evaporation (this problem is usually more severe when the medium is heated to 37 degrees Celsius). Therefore, special care must be taken when assembling cells into chambers and when changing culture medium if any evaporation has occurred (cells are very sensitive to rapid changes in osmolarity). In addition, during the imaging experiment, evaporation should be minimized either by using a sealed system, by covering the medium in an open chamber with an oil that has a lower density than water (usually mineral oil), or by humidifying the chamber during imaging. Note that, in general, the micro-environment provided by the small volume in a live-cell imaging chamber is inherently less stable than a carbon dioxide incubator and requires considerably more attention in every detail.
Choosing Cell Lines for Live-Cell Imaging
The choice of cell line used for live-cell imaging experiments is often dictated (and limited) by a number of factors, including the target biological observations of the investigation, the ability of the cells to be labeled with synthetic fluorophores, transfection efficiency, and the tolerance of a particular cell line to the rigorous culture chamber environmental and illumination conditions. All too often, a cell line that displays excellent properties in one of more of these categories either performs marginally or fails completely in another. For example, normal bovine pulmonary artery endothelial cells (BPAE line) can be fixed and stained using synthetic fluorophores to reveal intricate cellular structural details with exquisite clarity, but the line can only be transfected with fluorescent protein vectors at low efficiency (less than 5 percent) and is relatively intolerant to long-term illumination at low light levels that do not affect many other cell lines. Alternatively, rabbit kidney epithelial cells (RK-13 line) can be transfected at high efficiency with a variety of plasmids and are very tolerant to high illumination levels (including laser light) during time-lapse sequences extending for several days, but are not adequately stained with many of the common synthetic fluorophores (such as MitoTrackers) designed for live-cell imaging.
An overriding factor in the successful observation of biological phenomena in living cells is that the particular line chosen for study and imaging must display the necessary morphological and physiological properties to clearly demonstrate the concept of interest. In studies targeting mitosis, for example, many cell lines are less than adequate for imaging due to the fact that dividing cells become spherical and may detach from the substrate. Instead, cell lines that remain relatively flat and attached to the substrate during mitosis are far superior in revealing the fine details of the mitotic spindle during cell division. Among the most useful lines for mitosis studies are rat kangaroo kidney cells (PtK1 and PtK2 lines see Table 2), which can only be transfected at relatively low efficiency, but contain a small number of chromosomes that are easier to distinguish in the microscope. Several other kidney cell lines, including one from the pig (LLC-PK1) and another from the African green monkey (BS-C-1) also remain attached during mitosis and are far more susceptible to transfection. Both the pig and monkey cells contain more chromosomes than the rat kangaroo, but their ease of transfection and imaging make them excellent alternatives for investigations of mitosis. In addition to being useful for observations of the mitotic spindle, cells that remain flattened on the culture chamber glass during cell division can also reveal the distribution of other cellular components, such as the Golgi apparatus, cytoskeletal elements, endoplasmic reticulum, and mitochondria.
Table 2 - Useful Mammalian Cell Lines for Live-Cell Imaging Experiments
|Cell Line||Cell Type (Morphology)||Origin (Tissue)||Species||Marker (Applications)|
|B-16||Spindle||Melanoma||Mouse||Produces Melanin Cytoskeleton Investigations|
|BHK-21||Fibroblast||Kidney||Hamster||Plasmid and Virus Transfection|
|CHO-K1||Epithelial||Ovary||Hamster||Requires Proline DNA Transfection|
|COS-7||Fibroblast||Kidney||Monkey||T Antigen DNA Transfection|
|LLC-PK1||Epithelial||Kidney||Pig||Mitosis DNA Transfection|
|MDCK||Epithelial||Kidney||Dog||Cytokeratin Domes, Transport|
|PC-12||Aggregate||Adrenal Gland||Rat||Nerve Growth Factor Response|
|PtK2||Epithelial||Kidney||Rat Kangaroo||Cytokeratin DNA Transfection|
Investigations of the cytoskeleton should be conducted with cells that exhibit high levels of expression and specific localization of the protein(s) that are being studied. Filamentous actin stress fibers are usually much more clearly defined in fibroblast cells than epithelial cells, but there are many exceptions. Cytokeratin intermediate filaments form extensive networks throughout the cytoplasm, which are readily visualized with immunofluorescence or fluorescent proteins in several epithelial cell lines (although not universally). Unfortunately, cytokeratin networks are either poorly defined or virtually absent in fibroblast cells, as well as many varieties of epithelial and endothelial cells. Likewise, vimentin, desmin, peripherin, neurofilaments, lamins, and glial fibrillary acidic protein (GFAP) intermediate filaments form prominent structural networks in many cell types, but are difficult to detect in others. In almost all cases, the target cell line should first be fixed and tested with synthetic fluorophores and/or antibody labeling prior to attempting to localize fluorescent proteins for live-cell imaging.
A variety of applications coupling fluorescent proteins to live-cell imaging have opened many new avenues in the quest for information concerning dynamic processes in cell biology. The advanced fluorescence microscopy techniques of recovery after photobleaching (FRAP), resonance energy transfer (FRET), correlation spectroscopy (FCS), and speckle microscopy (FSM) have benefited significantly in their development from the use of fluorescent proteins. These ubiquitous molecules have also been genetically modified to produce a new generation of optical highlighters that can be photoactivated to specifically label individual members of a larger molecular population. Going a step farther, the coupling of Förster resonance energy transfer techniques to fluorescent proteins has yielded a new class of physiological biosensor probes that are useful for reporting various ions, such as calcium, sodium, potassium, chloride, and pH, in addition to a plethora of cellular events including enzymatic activity, changes in membrane potential, neurotransmitter release, and oxidation-reduction. The foundation for all of these powerful new techniques is built upon the imaging of living cells that are expressing genetically-encoded fluorescent probes.
Presented in Table 2 is a listing of several mammalian cell lines that have been of significant service to many of the live-cell imaging experiments reported in the scientific literature. The well-studied human cervical carcinoma (HeLa) line is an immortalized epithelial cell from which a wealth of information has been gathered. This hearty cell line can be transfected at high efficiency with most of the fluorescent protein vectors to produce high levels of expression and defined localization. Transformed African green monkey kidney cells (COS-7 line) have been used to explore protein dynamics in the Golgi apparatus and endoplasmic reticulum, whereas the hamster cell lines (BHK-21 and CHO-K1) are favorites for investigations involving the molecular and cellular biology of viruses, intracellular enzymatic activity, receptors, promoter function, and transport mechanisms. The other cells listed in Table 2 are also responsive to transfection, microinjection techniques, and labeling with synthetic fluorophores for long-term imaging experiments in widefield and confocal fluorescence microscopy.
Brief Overview of Live-Cell Imaging Chambers
Specimen chambers are an integral branch in the history of microscopy and a number of designs have been published over the years describing systems that offer excellent optical properties while allowing specimens to be maintained for varying amounts of time. Short term imaging experiments (20 to 30 minutes or less) can be conducted simply by attaching a coverslip containing adherent cells onto a microscope slide using spacers to keep the cells from being damaged (physical stress can induce autofluorescence in some cell lines). The coverslip can be secured with any one of a number of sealants, including molten agarose, rubber cement, vacuum grease, or a useful preparation known as VALAP (a 1:1:1 mixture of Vaseline, lanolin, and paraffin), to provide a watertight seal and eliminate evaporation of the culture medium. Thin gaskets cut from silicone rubber (also commercially available) or broken pieces of coverslip can be used as spacers to keep the cells from coming in direct contact with the microscope slide. Make certain the coverslip surface containing the cells is placed face down on the spacer, and fill the void between the coverslip and slide with a physiological buffer (such as phosphate buffered saline PBS). Seal the edges around the coverslip using the reagent of choice and place the microscope slide on the stage for imaging. Without growth medium and temperature control, the cells will function normally for only a few minutes, but this is often enough time to obtain the necessary images.
For longer term experiments, specially designed environmental chambers provide a mechanism for viewing and imaging living cells on the microscope stage, as well as keeping the culture very close to the optimum growth conditions for extended periods of time. In general, imaging chambers include a glass window, usually the thickness of a coverslip (approximately 170 micrometers), through which the cells can be readily viewed with objectives operating at high numerical aperture. Temperature control, a critical parameter for most cells, is often achieved using peripheral sources of infrared radiation or heated air (such as a hair dryer or egg warmer), a metal heating plate under thermistor control coupled directly to the chamber, or with optically transparent thin coatings of electrically conductive metal oxides applied by evaporation onto the coverslip surface to provide a more efficient conductive heat transfer to the chamber.
The wide variety of commercially available chambers that can be purchased (or, alternatively, easily constructed in-house) for imaging living cells generally fall into two basic functional categories: open chambers, similar to Petri dishes, which have free access to the atmosphere and closed chambers that are sealed to protect cells from evaporation of the culture medium. An open chamber system will usually allow quick access to the growing cells, thus readily permitting microinjection, addition of drugs, changing of the culture medium, or other manipulations to the cells. In contrast, closed chambers provide better insulation from the external environment, but make access to the cells more difficult. Most closed chamber designs include ports that permit the addition of fresh medium and drugs during the experiment without interrupting an imaging sequence. In these systems, perfusion is regulated by either a peristaltic pump, motor-driven syringe, or through a gravity-controlled manifold. When new solutions are added to a closed chamber, it is critical that before addition they are equilibrated to the same temperature as the cells. Furthermore, many cells are sensitive to shear, so perfusion of adherent cells attached to a coverslip should be performed at very low flow rates. Several of the more advanced closed chamber systems are designed to offer control of shear forces.
Figure 2 - Live-Cell Imaging Chamber and Controlled Environment Microscope Incubator
Many of the simplest commercial open chamber imaging systems are constructed by mounting a coverslip onto the bottom of a ordinary tissue culture vessel or Petri dish. Standard 35 and 60 millimeter sterile Petri dishes are available that have a small hole (approximately one centimeter in diameter) drilled in the dish with a 170 micrometer coverslip fused to the plastic to enable high-resolution imaging. Rectangular coverslips and microscope slides with a small single or multiple well plastic imaging chamber sealed to the glass are also commercially available, but are quite expensive. Both of these chamber designs are relatively simple to use, but they are not tightly sealed, so the amount of culture medium that evaporates over the course of an experiment must be carefully monitored. In addition, most of the simple imaging chambers do not include any heating system, and must be mounted on a microscope stage equipped with an auxiliary heating unit designed specifically to house the chamber. Without temperature control, simple open chamber system performance is only marginally better than using the sealed coverslip method described above.
Sealed closed chambers similar to the Bioptechs FCS2 live-cell imaging chamber illustrated in Figure 2(a) are more expensive than most of the simple open chamber systems, but they offer a far more controlled environment and can maintain cells in a healthy state for many hours (and even days or weeks). A typical closed chamber system provides two optical surfaces separated by a perfusion ring sealed with gaskets. This sandwich is then clamped together with a metal or composite housing that is designed to provide temperature control and secure adaptation to the microscope stage. With such an enclosure, the perfusion rate, media volume, temperature, atmosphere, flow geometry, and optical stability of the imaging chamber are controlled to a relatively high degree compared to open system chambers. Advanced closed chamber systems (Figure 2(a)) reduce the fluid exchange time, provide flow control of the perfusion medium in order to avoid disturbing adherent cells, offer superior temperature regulation, and maintain close proximity of the optical surfaces for observation using high numerical aperture microscope objectives. In addition, the user can define culture medium flow conditions across the cell surface to meet the experimental requirements. A wide variety of closed chamber live-cell imaging systems are commercially available.
The pinnacle of live cell imaging chambers effectively combines a cell culture incubator with an inverted microscope to provide almost total control of the environment, an example of which is presented in Figure 2(b). The incubator enclosure is most often constructed of Plexiglas and surrounds the microscope stage, objectives, fluorescence filters, and transmitted light condenser. These chambers can be used with a variety of culture vessels, including standard culture bottles, Petri dishes, microscope slides with mounted coverslips, and many of the other open and closed systems discussed above. Temperature is maintained with an external heating unit (usually forced air) and the carbon dioxide concentration is controlled with a sensing unit coupled to a regulator that is fed by a cylinder of pure gas. These units can also be equipped with humidity control and several designs provide rubber glove access to avoid disturbing the environmental equilibrium when manipulating the cells during imaging. In order to maintain a high degree of temperature control, several of the more sophisticated incubator chambers enclose virtually the entire microscope with the exception of the eyepieces, camera, and lamphouses. On the downside, environmental chambers can impede rapid access to the specimen and are cumbersome when repeated manipulation is necessary. In addition, the high humidity level inside the chamber can add to the expense of maintaining the instrument due to premature degradation of gear lubricants and the oxidation of metal surfaces and lens coatings. Most of the microscope manufacturers offer a custom incubator option for their inverted microscopes, while aftermarket suppliers fill the gaps with both simpler and more advanced models, as well as a host of useful accessories.
As discussed above, a wide spectrum of open and closed live-cell imaging chamber designs are commercially available, many of which are intended for quite specific applications. It is worthwhile to investigate the various options available when embarking down the long road to successful live-cell imaging. Most investigators have favored techniques that reflect their experience using a particular system, and these preferences span the gamut of chamber designs. In fact, several functional systems have been reported that were constructed with readily available home repair insulation sheeting, duct tape, and low-cost humidifiers connected to the chamber using clothes dryer exhaust tubing. The key point is that in a given experiment, the chamber must maintain an optimal environment for cellular function and provide a clear optical window in which to capture the events occurring within the chamber. Because the experimental variables differ from one investigation to another, design preferences can change and the best approach is to test a variety of systems in order to identify the one that is best suited to the cell line and the experimental conditions.
With all live-cell imaging configurations, difficulties can arise when experiments are performed on preparations that require temperatures significantly different from that of the surrounding laboratory environment (note that temperature fluctuations in live-cell imaging are usually the rule rather than the exception). Cellular function is extremely sensitive to temperature variations, with changes of even a couple of degrees having profound effects on cell physiology. A variety of methods are available for controlling the temperature of cells on the microscope stage, and many of the commercial systems described above include heating elements directly coupled to the chamber. Although this strategy provides a simple integrated solution, temperature control is too often limited to the chamber itself and ignores associated components that might exert negative influences on maintaining a steady temperature. Among the most significant factors associated with temperature fluctuations is that the microscope stage, frame, and objectives can act as heat sinks and counteract the efforts of the specimen heating system. This problem is compounded when immersion objectives are used because the optical coupling medium, which can be oil, glycerin, or water, has a much higher thermal conductivity than air. Coupled with the close proximity of high numerical aperture objectives to the specimen, as well as the thermal load of the objective itself, the entire system can be rapidly deprived of heat if the objective is not thermally controlled. In the case of the static chambers described above, the area directly under the objective is often up to 5 degrees (Celsius) cooler than the remainder of the specimen chamber.
Figure 3 - Objective Heater Designs for Live-Cell Imaging
Depending upon the configuration, the entire microscope can be enclosed and heated, but several critical temperature control issues can be avoided simply by using commercially available objective heaters (see Figure 3) that employ circulating heated water or resistive heating elements. Objective heaters, when combined with a suitable specimen-heating system, can partially offset the temperature gradient between the specimen and the front lens elements. Note, however, that even with an objective heater, there can still be a temperature gradient along the objective barrel or between the objective and the microscope itself. The adjustable heating blanket and circulating water jacket objective heaters illustrated in Figures 3(a) and 3(b) are essentially the same from a thermodynamic point of view. The only difference is that heat is electrically produced in the blanket (Figure 3(a)), while heat is externally generated and transferred by a fluid into the jacketed objective (Figure 3(b)). In both cases, heat transfer is relatively inefficient with much of the heat being radiated away from the objective to the region directly beneath the specimen instead of being transferred into the objective. The unfortunate result is excessive external heat being convectively channeled upwards in the vicinity of the specimen to produce large temperature variations and excessive thermal cycling in the chamber control system.
Microscope objectives have thermal profiles that vary as a function of their physical parameters. Most objectives are designed and sold for the purpose of fixed-cell microscopy (conducted at room temperature), so when selecting an objective to be employed in live-cell imaging, care should be taken to consider only those objectives with the ability to be efficiently heated. Aside from objective heating issues, cycling of the heating system can alter the coverslip position and cause the specimen to drift out of focus. In addition, the investigator should be aware that repeated heating and cooling of the objective has been incorrectly reported to considerably shorten the lifespan, especially in terms of the strain-free character of internal lens elements. In fact, there is little reliable evidence that temperature cycles affect the strain characteristics in specialized objectives, and most can withstand temperatures up to 50 degrees Celsius without damage. The only negative effect of heating microscope objectives is that the retraction stopper barrel lubricant can increase in viscosity (achieving the pliability of gum) over a shorter period of time than with objectives that are not heated. However, in the case of heated immersion objectives, the tendency of immersion oil to creep into the barrel often prolongs the lifetime of the original lubricants.
For permanent installations, a large box can be built around the microscope and heated with warm air. In this case, most of the microscope can be equilibrated to a single temperature to provide the advantage of eliminating any movement resulting from thermal expansion of the microscope components. However, air currents surrounding the specimen chamber itself must also be minimized. When constructing an enclosure, access to the microscope and its adjustable components may be limited, so it may be worthwhile to construct the box from relatively common and inexpensive components in case significant modifications are required.
A final consideration is tight control of temperature not only for the microscope, but also the entire laboratory. Modern microscopes are fabricated with a wide spectrum of materials, including aluminum, plastic, composites, glass, brass, and steel, all of which have different thermal expansion coefficients. Even a change of a single degree Celsius can produce unwanted movements in the microscope optical train, resulting in focus or alignment shifts. Air conditioning or heating ducts that are in close proximity to the microscope will often produce localized temperature fluctuations that ultimately result in focus problems. For long-term observation, many investigators build a large thermostatically controlled box around the stage (see the discussion above) or even place the entire microscope in a room that is maintained at 37 degrees Celsius (a relatively uncomfortable working situation). The exact strategy that is employed ultimately depends on the specific application, but it is absolutely critical to consider these issues when designing a live-cell imaging system and the laboratory in which it will be housed.
Laboratory Environmental Considerations
When choosing a room that will be utilized for live-cell imaging experiments, it is necessary to ensure that adequate ventilation is available to dissipate the ozone released by mercury and xenon arc-discharge lamps, as well as the fumes from organic solvents used to clean optical surfaces and disinfect the microscope stage. Allow sufficient space around the microscope system for proper ventilation as well as cleaning of the floors, benches, tables, and experimental apparatus. Equipment failures can often be traced to air intakes that are clogged, located close to the floor, or placed in an inaccessible location. The laboratory should be kept meticulously clean and maintained in an orderly fashion to reduce the levels of dust, smoke, and other damaging vapors that can diminish optical as well as electronic performance. In order to reduce the incidence of live-cell culture contamination by microorganisms, the microscope stage and surrounding area should be periodically wiped with 70-percent ethanol or commercial antiseptic towels. Culture media spills, an unavoidable factor in live-cell chamber manipulation, should be cleaned immediately and the surrounding area thoroughly disinfected.
Among the mechanical vibration sources that can affect microscope performance are central heating and cooling units (or air handlers) in the attic or on the roof of the building, refrigerators, low-temperature incubators (even in adjacent rooms), and traffic through nearby hallways. Vibrations from refrigerators and other sources that may not be immediately obvious can have a severe impact on microscope stability. A variety of techniques can be applied to reduce room and building low-frequency vibrations, including feedback-controlled isolation tables that are gas-filled (the most expensive option) and relatively low-cost flexible synthetic polymer vibration isolation pads (see Figure 4). The latter are marketed in a variety of geometries and damping levels to suit a wide spectrum of configurations. A combination of the vibration pads and a heavy sheet of half-inch aluminum or a pre-drilled isolation platform often will reduce vibrations to an unnoticeable level. Although inverted microscope frames are much heavier and usually less sensitive to vibration compared with upright microscopes, they still benefit from isolation. High-frequency vibrations can, in many cases, be substantially reduced by loading the table top with additional mass, such as lead bricks. Finally, when very small displacements of sub-resolution features are the subject of investigation, additional steps may be required, such as working very late at night or early in the morning, when the environment is quieter.
Figure 4 - Vibration Isolation Systems
In addition to vibrations conducted through the floor, possible airborne sources must also be considered. Equipment cooling fans or air-conditioning fans within the laboratory, and even exhaust noise from nearby vehicles (conducted through laboratory windowpanes), have been reported to produce troublesome vibrations for high-resolution microscopy on otherwise stable equipment. The vibration of cooling fans for lasers and other major equipment can also affect live-cell imaging stations. A section of flexible duct tubing placed between the cooling fan and the laser head can be effective in decoupling fan vibrations. In perfusion and gravity-fed media systems, vibrations due to gas bubbling through the feeding solution can couple through the tubing to the specimen chamber and produce periodic mechanical displacements. Acoustic noise can also be a source of vibration and reduced microscope stability. Noisy fan units should be replaced and other equipment that tends to produce excessive amounts of noise (and possibly vibration) should be relocated, if feasible, to another room. Microscopes are often equipped with built-in light sources and power transformers that can gradually heat the microscope frame and produce a slow focal drift of the specimen.
Control of room temperature for an otherwise stable microscope live-cell imaging system is another major concern and can be one of the most significant problems encountered when recording time-lapse sequences. A microscope that is confined to a small and poorly ventilated room can produce a considerable increase in ambient temperature. The heat load resulting from light sources, temperature controllers, shutters, cameras, computers, and other equipment may exceed the nominal capacity of the room, requiring the services of an auxiliary cooling system. Additional cooling improves the operation of all electronic equipment provided it does not introduce additional vibration and dust. In this regard, the cooling-fan capacity of microscope system computers and high-speed disk drives may become insufficient as the chassis is filled with heat-producing cards, such as those used for camera controllers, image processing software, and additional memory. Problems are manifested as system crashes, increased noise, and, in the worst extreme, lost data. Addition of a low-noise box fan to the computer housing can often alleviate this difficulty.
Although modern inverted microscopes (those most favored for live-cell imaging experiments), such as the Nikon Ti2 series, are designed to be solid stand-alone instruments through a considerable amount of engineering effort to minimize vibration sources, aftermarket auxiliary components can often compromise this inherent stability. Fast shutters and optical filter changers (filter wheels) produce significant levels of vibration during operation and, when attached directly to the microscope frame, often induce perturbations that can last for tens to hundreds of milliseconds. In addition, many research-level microscopes often contain built-in motorized components (nosepieces, focus controls, etc.) that can be sources of vibration unless the manufacturers limit the speed of such devices and/or introduce delays between motorized functions and image acquisition. Vibration from the auxiliary components can be dramatically reduced or even eliminated by mounting these devices independently on separate stands adjacent to the microscope. Rigid aluminum optical breadboards containing pre-drilled mounting holes are commercially available, and are ideal for housing both the microscope and accessory components.
Fluctuations in the axial position of the microscope focal plane during the collection of sequential images from living cells is one of the most serious and frequently encountered problems in time-lapse microscopy. Often termed focus drift, changes to the microscope focal plane usually occur due to temperature variations in the imaging chamber or within the room in which the instrument is housed. Generalized qualitative analysis of the relationship between the thermal state of a microscope and the focal plane position indicates that a one-degree (Celsius) increase or decrease in temperature can shift the focus by approximately one-half micrometer (500 nanometers). Irregularity of the surface in glass coverslips, as well as mechanical instability arising from gear slippage, compression of lubricating grease layers, and settling between the moving components of the microscope, although not usually a major source of concern, can also contribute to focus drift. Without a feedback device to continuously monitor and correct focus, one of the best remedies for drift is to employ a thermostatically controlled enclosure that fully envelops the microscope and associated components. Although it is possible to obtain image sequences that remain in focus for short periods of time without auxiliary equipment, a high percentage of longer term experiments fail due to focus drift. In almost all cases, thermal instability leading to flexing of the coverslip or temperature gradients in the microscope optical train can usually be traced as the source of the failure.
Defeating focus drift should be a principal consideration during configuration of a microscope for time-lapse imaging, especially when using high numerical aperture objectives where the narrow depth of focus (approximately 300 nanometers) requires a focal position that is held to within a 100 nanometers of the initial plane. The entire system, including the microscope, camera, shutters, filter wheels, illuminators, live-cell chamber assembly, and host computer should be brought to operating temperature for at least 24 to 48 hours prior to initiating time-lapse imaging sequences. When assembling the imaging chamber, ensure that coverslips containing adherent cells are mounted securely in their housings and that the chamber itself is positioned on the stage in a manner that does not allow movement either in the lateral or axial directions. At high imaging resolutions, oil immersion objectives can be the source of focus drift as the oil spreads across the chamber coverslip and the front lens element (dry objectives do not have this problem). An objective heater, discussed above, should be used for all configurations that employ immersion objectives. Once the microscope is at the correct operating temperature and all other equipment is staged for the experiment, monitoring focus with a preliminary time-lapse sequence over a period of 12 to 24 hours using a fixed and permanently mounted specimen will provide an excellent indication of the system stability.
A variety of commercially available software and hardware solutions have been introduced by microscope and aftermarket manufacturers to contend with focus drift. Several of the hardware devices are autofocus systems that measure the distance between the objective front lens and the specimen by sensing light or sound reflected from the lower surface of coverslip (closest surface to the objective), such as the Nikon Perfect Focus System 4 for Ti2 inverted microscopes. This approach can be hampered, however, when high resolution oil immersion objectives are used, due to loss of contrast and reflectivity as the sensing light passes through the oil. The most advanced autofocus systems use low intensity near-infrared laser or light emitting diode (LED) sources to reflect a beam of illumination through the objective and onto the upper surface of the coverslip (supporting the cells and bathed by the culture medium), subsequently recovering the reflected light with the objective and passing it on to a detector that controls a feedback circuit to adjust the position of the objective in relation to the coverslip-culture medium interface.
Figure 5 - Inverted Microscope Automatic Focus System
Instantaneous feedback of specimen position enables the distance between the coverslip and objective to be detected in millisecond intervals during observation, thus automatically correcting for focus drift in real time. These systems are especially useful in terminating focus drift resulting from a temperature drop that occurs when changing and perfusing culture medium or adding reagents (such as drugs) to the culture. In addition, examining the cells to determine candidates for imaging is facilitated by constant focus corrections while translating the stage during observation and setting locations for multipoint image collection. Software accompanying the autofocus systems allows freely selectable focal planes through adjustment of an offset control. Due to the long wavelengths used by the laser or LED light source, the near-infrared detection system does not intrude on wavelengths used for observation and should be invisible to the fluorescence detector.
Illustrated in Figure 5 is the schematic diagram of a typical automatic focus drift correction mechanism for an inverted microscope (Figure 5(a)) along with a spectral window comparing fluorescence emission profiles of common fluorescent proteins with the wavelength of the autofocus laser spectral line (Figure 5(b)). The laser (or light emitting diode) is focused into the rear focal plane of the objective and introduced parallel to its centerline axis with a specified offset. The positioning feedback loop corrects for minute geometrical shifts due to focus drift in the position of the beam that is reflected from the interface between the upper coverslip surface and the culture medium. The spectral emission profiles outlined in Figure 5(b) represent several of the most useful fluorescent proteins covering the wavelength region between 450 and 700 nanometers, including enhanced cyan (ECFP), green (EGFP), and yellow (EYFP) fluorescent proteins, as well as reef coral red-shifted variants, monomeric Kusabira Orange (mKO) and mCherry. The 870-nanometer laser line used by the focus drift correction system does not interfere with the fluorescent protein signals.
In laser scanning confocal microscopy focus drift correction during time-lapse investigations is often accomplished by obtaining multiple axial images at each time point with subsequent analysis of every image stack to identify a common focal plane that corresponds to a pre-selected reference point. Other approaches employ software algorithms that determine the focal position between time points using contrast functions that record images in successive upward and downward steps and compare the results until the highest level of contrast is obtained (at which point the focus is set for that interval). Software techniques rely on a relatively constant level of specimen contrast, however, which is often not the case with live-cell imaging where debris and other artifacts can randomly float into the viewfield and alter the apparent optimum focal plane.
Phototoxicity and Photodamage
Aside from the toxicity that occurs due to excessive concentrations of synthetic fluorophores and over-expression of fluorescent proteins, the health and longevity of optimally labeled cells in microscope imaging chambers can also suffer from a number of other deleterious factors. Foremost among these is the light-induced damage (phototoxicity) that occurs upon repeated exposure of fluorescently labeled cells to illumination from lasers and high-intensity arc-discharge lamps. In their excited state, fluorescent molecules tend to react with molecular oxygen to produce free radicals that can damage subcellular components and compromise the entire cell. In addition, several reports have suggested that particular constituents of standard culture media, including the vitamin riboflavin and the amino acid tryptophan, may also contribute to adverse light-induced effects on cultured cells. Fluorescent proteins, due to the fact that their fluorophores are buried deep within a protective polypeptide envelope, are generally not phototoxic to cells. However, many of the synthetic fluorophores, such as the MitoTracker and nuclear stains (Hoechst, SYTO cyanine dyes, and DRAQ5), can be highly toxic to cells when illuminated for even relatively short periods of time. In designing experiments, fluorophores that exhibit the longest excitation wavelengths possible should be chosen in order to minimize damage to cells by short wavelength illumination.
Presented in Figure 6 are three examples of phototoxicity induced by the illumination of cells labeled with synthetic fluorophores or expressing fluorescent fusion proteins. The normal rabbit kidney cells (RK-13) shown in Figure 6(a) were first transfected with a fusion vector of EGFP and the SV40 virus T-antigen nuclear targeting sequence to localize green fluorescence within the nucleus. The cells were subsequently treated for 20 minutes with MitoTracker Red CMXRos and imaged sequentially in 30-second intervals for 24 hours. After several hours, vacuoles began to form in the region containing mitochondria surrounding the nucleus. Illustrated in Figure 6(b) is a single human glioblastoma cell (U-251 line) expressing mCherry fluorescent protein fused to human beta-actin after several hours of observation. Note the significant deterioration of cellular structure that has occurred as the cell appears to enter necrosis. Detachment of Swiss albino mouse embryo cells (3T3 line) after an hour of imaging following treatment with the DNA-binding nuclear stain Hoechst 33342 is depicted in Figure 6(c). Ultraviolet-absorbing nuclear dyes (such as Hoechst) usually demonstrate phototoxic effects more quickly than do probes that are excited at longer wavelengths in the visible spectrum.
As discussed above, the synthetic biological buffer HEPES has been reported to produce a phototoxic effect on cells in some circumstances (although in many cases this might be attributed more directly to inadequate levels of bicarbonate). The investigator should keep in mind that all cells are intrinsically photosensitive, and adding fluorophores or aromatic components to the culture medium only compounds this sensitivity. Damage by the free radicals generated through excited state fluorophores can only be limited, not prevented. However, healthy cells have inherent enzymatic mechanisms for converting free radicals to less harmful compounds and can tolerate fluorescence excitation provided their enzyme systems are not saturated. Reducing the level of oxygen in the medium, provided the cells can survive oxygen withdrawal, can serve the dual purpose of limiting the degree of photobleaching and free radical production.
Figure 6 - Cellular Phytotoxic Effects from Synthetic and Genetic Fluorophores
Regardless of the potential for cells to enzymatically deal with toxicity arising from fluorophores and culture medium components, the exposure of cells to light in the imaging setup should be reduced to the lowest possible level in order to limit other sources of cell damage and to minimize the potential for artifacts in the experiment. The illumination dose per image can be limited by the judicious application of neutral density filters (for arc-discharge lamps), lowering the output power of the illumination source (for lasers), and programming shutter assemblies to restrict exposure to fluorescence excitation light only during image capture. In addition, fluorescence filter bandwidths should be carefully chosen to reduce the number and intensity of unnecessary wavelengths that will illuminate the cells with light that is not useful for imaging. The ability to successfully decrease illumination levels in live-cell imaging is aided by using high numerical aperture objectives that feature the best light throughput and the fewest optical elements.
Even in the absence of fluorophores, the sensitivity of mammalian cells to ultraviolet light exposure has been well documented (a phenomenon known as photodamage), and many cell lines are at least equally sensitive to blue light in the wavelength regions used to excite cyan and green fluorescent proteins. To minimize photodamage and phototoxicity when setting up an imaging experiment, visualization of the living cells during microscope configuration must be performed at the lowest light levels under which the cells can be observed, and this should be conducted as quickly as possible. Neutral density filters (or very low laser power) should be used to attenuate the illumination source and visualization of the cells is best accomplished with the digital imaging system. Observing the cells through the microscope eyepieces takes several seconds, which is at least tenfold longer than is often required to obtain an image of sufficient quality for cell selection and focusing. Alternatively, candidate cells for imaging can be located using brightfield, phase contrast, or differential interference contrast (DIC) imaging modes. In all cases, a green or red interference filter should be used during setup to improve contrast and minimize exposure of the cells to blue light.
The widespread application of 546-nanometer (green) filters for live-cell imaging studies originated because this wavelength region matches one of the major spectral lines emitted by mercury arc-discharge lamps. However, achieving adequate levels of illumination intensity is rarely a problem with live-cell investigations and modern objectives are corrected to such a degree that they do not require the use of green light for high-resolution imaging. Thus, the choice of illumination wavelengths should be limited to those in the regions best tolerated by the cells. Immediately prior to cell division (prophase), most mammalian cell lines are very sensitive to ultraviolet and infrared light, and are least sensitive to red, green, and blue light, in descending order. Thus, in order to minimize photodamage, red interference filters with a bandpass region between 600 and 650 nanometers are the ideal choice for live-cell observations. Autofluorescence in mammalian cells and tissues is also reduced in the longer wavelengths of the visible light spectrum. Note that optical resolution is dependent upon wavelength and red light yields the lowest theoretical values for resolution due to the longer wavelengths involved. In most cases, however, the use of red light is not the limiting factor because the degree of resolution achieved in live-cell imaging is often compromised by internal cellular motion, temperature drifts, imperfections in the optical system, and illumination fluctuations.
In choosing filters for live-cell imaging experiments, bandwidths should be carefully selected so that even trace levels of infrared and ultraviolet light are eliminated. Even though modern bandpass filter designs perform well in the central regions of the visible light spectrum, they often pass radiation at very low and very high wavelengths. It is therefore advisable to install specialized glass filters near the illumination source(s) to block damaging ultraviolet and infrared wavelengths. Mercury and, to a lesser extent, xenon lamps produce high levels of ultraviolet light, while tungsten-halogen (transmitted) lamps emit significant amounts of infrared light. As a final step, electronic shutters should be installed (for both tungsten-halogen and arc-discharge lamps) to limit exposure of the cells to damaging radiation during periods when images are not being captured. Judicious shuttering of the illumination source is one of the most important factors in successful live-cell imaging experiments.
Advances in detector technology over the past few years have made it feasible to further reduce illumination levels in live-cell imaging experiments. Increasingly sensitive photomultiplier tube cathodes for confocal microscopy and advanced charge-coupled device (CCD) camera systems for widefield microscopy are continually being introduced. The intensified and electron multiplying camera systems now available are capable of imaging living cells with high sensitivity at light levels that are exceedingly low. Many of these cameras employ back-thinned CCDs, which are usually cooled and feature high quantum efficiency across the visible and near-infrared spectral regions to further increase sensitivity. If the best cameras are not available, sensitivity can be increased by combining the signal from multiple pixels (a process known as binning) at the expense of spatial resolution. In confocal microscopy, maintaining low zoom ratios will reduce the level of phototoxicity to cells. Increasing the confocal zoom factor causes the total amount of laser light to be scanned over a smaller region of the specimen, thus exposing the cells to more intense illumination.
Choosing the exact level of light attenuation and the correct exposure time is almost always an empirical exercise. For a new cell line with unknown parameters, the best strategy is to attenuate the light as much as possible and apply very short exposure times so that subcellular structures are just barely visible in the acquired image. A good place to start is a neutral density filter with an optical density of 1.0 and an exposure time of 100 milliseconds or less. For laser scanning confocal microscopes, start with a laser power of approximately 1 percent and increase the voltage (and gain, if necessary) of the photomultiplier. Use pixel dwell times about 50 percent shorter than those that usually produce adequate signal levels. If the cells are able to tolerate this light level through a long-term time-lapse experiment, then the illumination intensity and exposure time can slowly be increased in subsequent experiments until a workable compromise is achieved between signal-to-noise and cell viability. It should be noted that a nonlinear relationship often exists between the total amount of light exposure a cell can tolerate and the length of individual exposure times. In general, cells appear to be the healthiest when exposed to very brief pulses of light, since extended exposures (greater than one-half second) are often lethal over long periods of time.
Monitoring Cell Viability and Variability
After the live-cell imaging chamber has been loaded with fresh cells, assembled, and mounted on the microscope stage, the next step is to visualize the cells to establish their overall condition and morphology, and to identify candidates that are appropriate for imaging. In a vast majority of experiments, especially when imaging cells that have recently been transiently transfected with fluorescent proteins, there will be a significant amount of morphological variability in the cell population. It is not uncommon to observe cells that were either not transfected or exhibit poor localization of the probe. In addition, a percentage of the transfected cells will over-express the fluorescent protein, often to the point of creating a potentially toxic effect. Other cells may exhibit common health problems (as illustrated in Figure 7) manifested in the form of detachment from the substrate, excessive vacuole formation, swollen mitochondria, and cytoplasmic blebs. The same cell line in different cultures can demonstrate varying patterns of beading and blebbing that may be a symptom of dissimilar stress factors. Declining health often affects the rate of cell growth and motility, usually resulting in a general decrease in activity. However, poor health is not always indicated by a decline in cellular activity as compromised cells can exhibit a marked increase in Brownian-like organelle motions. Cells showing even a slight deviation from a normal and healthy appearance should not be pursued for imaging and data collection. Furthermore, if more than 50 percent of the population is judged as unhealthy, the entire culture should be discarded and exchanged for one that is in better physiological condition.
Figure 7 - Visual Symptoms of Morphological Variation in Unhealthy Cells
Many live-cell imaging experiments are performed with only a single or, at most, a few cells. The investigator should keep in mind that the morphology of cells in culture can be quite varied with regard to the apparent phenotypes present. This heterogeneity can be the result of cells in different phases of the growth cycle or possibly intrinsic differences among individual members of the population (the latter is more apparent in primary cultures). For this reason, it is often necessary to record data from a number of individual cells in order to gain a statistically significant sampling of cellular behavior and dynamics. As discussed above, the investigator should bear in mind that the imaging conditions in live-cell observations must be minimally perturbing to the cells in order not to significantly affect the experimental result. Among the most critical aspects of live-cell imaging experiments is establishing a criterion for judging the health of the cells under study so that the success of the experiment can be evaluated objectively. The exact criteria will vary depending upon the experiment, but one of the most important factors to be considered is whether the expected result was achieved without damage to the cells incurred by the imaging process. In some cases, the phenomenon under study can be matched to results obtained with fixed cells, but this is often not possible.
Imaging experiments should be monitored to ensure that the experimental conditions (culture medium, buffering strategy, atmosphere, chamber configuration) do not substantially alter the growth rate, mitotic index, or apoptosis properties of the cells. If possible, the potential negative effects of each experimental component should be separately assessed. For example, cells should be grown in the culture medium used for the imaging experiment in a standard carbon dioxide incubator and judged for viability, mitotic index, and general morphological variations. As a final check, the cells can then be installed in the environmental imaging chamber (without illumination) for a number of hours and then similarly assessed. This strategy isolates the various contributions to general cell health and readily identifies any procedures that require modification.
As discussed above, cells are exposed to high doses of illumination during live-cell imaging experiments and those that are labeled with fluorophores are potentially venerable to the generation of reactive molecular species that can seriously affect cellular function. It is therefore safe (and prudent) to assume that at least some level of damage has occurred during the imaging experiment and steps should be taken to determine whether it is significant enough to have produced detrimental effects. This is best accomplished by leaving the cells in the imaging chamber on the microscope stage after completion of the experiment with subsequent periodic examination to determine whether the cells initiate apoptosis or enter and complete mitosis. During the post-experiment evaluation process, time-lapse images can be recorded at widely spaced time intervals (10 to 30 minutes) over a number of hours.
The criteria for determining cell viability depend on the appropriate pathways present in the cells under examination. Some transformed mammalian cell lines have defunct cell cycle checkpoints that render determinations of mitotic and apoptotic progression useless. Each cell type should be carefully evaluated to determine the criteria that can be used for viability assessment following imaging sequences with the microscope. Among the simplest assays following an imaging experiment is to compare the apparent health and morphology of the cells that have been exposed to light during the investigation with neighboring cells in the same chamber that haven't been subjected to illumination. Similar features in the two populations, when observed with phase contrast or DIC, are a good indication that general health has not been compromised. This observation should be followed with fluorescence imaging to ascertain whether fluorophore localization has changed during the course of the experiment. Finally, both populations of cells should be intermittently observed for several hours (or days) following the experiment to see if the cells subjected to imaging behave similarly to the non-imaged cells. Comparisons of this type often reveal whether significant damage has occurred to the cells under study.
Examination of the mitotic pathway during live-cell imaging is an excellent mechanism with which to monitor cells for photodamage. Cells that are just beginning the mitotic cycle by initiating chromosome condensation (prophase) often fail to enter prometaphase and subsequently complete cell division when damaged by excessive illumination. Scanning the culture with the microscope operating in differential interference contrast mode can reveal those cells that are commencing mitosis, and a suitable candidate can be isolated for closer examination under the actual experimental imaging conditions. If the chromosomes undergo decondensation during the observation period and the cell fails to re-enter mitosis within several hours, there is a good likelihood that photodamage has occurred. Note that changing the culture medium will also initiate decondensation of prophase chromosomes in many cells, so this analysis should be performed several hours after the cells are placed on the microscope stage. Cell lines differ widely in their ability to tolerate light during mitosis. Established lines and primary cultures of human and animal cells can be extremely sensitive to light, whereas cells derived from embryos (such as fruit flies and nematodes) often lack pathways to arrest the division cycle in response to DNA damage and are more tolerant to photodamage.
Figure 8 - Microbial Contamination in Mammalian Cell Cultures
Aside from the numerous problems associated with photodamage and phototoxicity, as well as the rigorous maintenance requirements incumbent on live-cell imaging, the investigator must also be alert to the possibility of microbial contamination during the course of the experiment. The most common infections are those that occur due to bacteria, fungi, mycoplasma, yeasts, molds, and in rare circumstances, protozoa. Unless it becomes a frequent event, the nature of the infection or species involved is not as important as determining where the contamination originated. In general, rapidly growing microorganisms are less problematic due to the fact that they are usually readily detected and the culture can be quickly discarded. More important are those infections whose presence is cryptic, unable to be visualized during routine examination of the culture because of the physical size, or enabling the invader to grow at a level that escapes detection. Overuse of antibiotics in the culture medium is a common problem that often results in a low-level contamination, which can remain undetected for long periods of time and may ultimately interfere with normal mammalian cellular function.
Presented in Figure 8 are micrographs illustrating three of the most common visible sources of microbial contamination in live-cell cultures. Unchecked bacterial growth (Figure 8(a)) is readily detected at high magnification (often accompanied by a rapid drop in culture medium pH) and can even be visualized as turbidity in the culture medium with the unaided eye when the organism numbers begin to reach saturation. Yeast (Figure 8(b)) grow far more slowly than bacteria, but have a distinct colonial motif that reveals their presence. Mold contamination (Figure 8(c)) usually overtakes a culture within 24 to 48 hours, and can often be identified as a bushy, fibrous intrusion. Perhaps the most serious form of contamination, however, is not obvious using routine contrast-enhancing microscope techniques (phase contrast and DIC). Mycoplasma (not illustrated) can seriously alter cell behavior and metabolism, but are not easily identified in cell cultures other than through often subtle signs of gradual deterioration. Assays for mycoplasma should be routinely conducted to ensure cultures are free of this serious artifact in live-cell imaging. A variety of detection methods are useful for revealing the presence of mycoplasma, including fluorescent staining (using the nuclear dye Hoechst), polymerase chain reaction, and autoradiography.
Dynamic imaging of biological activity was introduced in 1909 by French doctorial student Jean Comandon, who presented the earliest reported time-lapse cinema films of syphilis-producing spirochaetes, 5 years before Charlie Chaplin made his first movie. Comandon's technique, which he called microcinematography, enabled the production of movies capturing events in the microscopic world that could be recorded using an enormous cinema camera bolted onto a fragile darkfield microscope. These films proved instrumental in teaching physicians how to distinguish disease-causing spirochaetes from those that are harmless, and demonstrated how time-lapse observations can be employed to gain important biological information without recourse to image analysis, processing, or even empirical quantitative measurements. For the next 75 years, many microscopists adapted increasingly advanced cinema film cameras to the microscope in order to generate progressively better films at much higher resolution.
As tube-based video cameras became affordable in the late 1970s and early 1980s researchers began to couple these devices with optical microscopes to produce analog time-lapse image sequences and real-time videos. The tube camera ultimately gave way to the area array CCD in the early 1990s, heralding a new era in photomicrography and signaling the ultimate demise of film. With the advanced digital camera systems available in the 21st century, the increasingly popular technique of time-lapse cinemicrography is becoming broadly applied to capturing events that occur in living cells over periods ranging from a few seconds to several weeks (or even months). The technique involves repeated imaging of a cell culture at defined time points, thereby providing information on myriad dynamic processes that often occur with a wide distribution of time scales. When time-lapse investigations are coupled to labeling cells with synthetic fluorophores and genetically encoded fluorescent proteins, events at the subcellular and molecular levels can be investigated.
Time-lapse imaging can be performed in two spatial dimensions using widefield techniques and extended even further to three-dimensional imaging with confocal microscopy. In addition, modern confocal microscopes are equipped with line-scanning software for rapid and repeated imaging of single scan lines. Two-dimensional time-lapse imaging involves sequential capture of single focal planes (x-y in widefield microscopy and x-y, x-z, and y-z in confocal microscopy), whereas three-dimensional imaging produces optical stacks from multiple focal planes in a variety of dimensional formats with thick specimens. When a single focal point in the lateral plane (x and y) is combined with z-stack imaging as a function of time, the technique is referred to a 4-D time-lapse imaging. Likewise, adding a fifth dimension (wavelength) yields 5-D imaging whereas adding either multiple wavelengths or multiple lateral regions to the 4-D stack is referred to as 6-D time-lapse imaging. As described above, the time intervals for sequential image gathering using these techniques can range from milliseconds to days or even months.
A growing number of small molecule, vital synthetic fluorescent probes that yield highly specific cellular or subcellular labeling patterns are now commercially available. In addition, the huge effort to produce useful fluorescent proteins having emission colors spanning the visible and near-infrared spectrum is beginning to produce encouraging results. Green fluorescent protein and related spectral variants are now routinely being fused to other proteins of interest to reveal details concerning protein geography, movement, lineage, and biochemistry in living cells. In this regard, these biological probes have provided an important new approach to understanding protein function, which is the next logical step for investigations of cellular processes now that the genome sequence of many organisms has been determined. The inherent brightness and photostability of many fluorescent proteins render them well-suited for the repeated imaging required in time-lapse studies. Together, the synthetic and genetically encoded fluorescent probes are affording a seemingly endless array of possibilities for imaging molecular components in living cells.
Figure 9 - Time-Lapse DIC and Fluorescence Live-Cell Imaging
Illustrated in Figure 9 are several images from a 24-hour time-lapse sequence of rabbit kidney epithelial (RK-13) cells that were captured using a combination of fluorescence and differential interference contrast. The cells were transfected with a chimera of mCherry fluorescent protein fused to human beta-actin to reveal the distribution of this cytoskeletal component in the main cell body as well as the lamellapodia. The white arrow in Figure 9(a) indicates the initiation of a cytoplasmic ruffle near the large pool of actin in the central region of the cell. As the sequence progresses, the small ruffle begins to swell and extrude towards the left-hand side of the image, in a wave-like motion, concentrating the brightly labeled actin fusion protein into the leading edge as it grows (indicated by the arrows in Figure 9(b) through 9(d)). As the lamellapodium spreads to cover a large area, small clusters of labeled actin form behind the leading edge (arrows in Figures 9(e) and 9(f)) in structural elements that may be podosomes involved in the process of cellular adhesion to the glass substrate.
Time-lapse imaging techniques are significantly aided by the application of critical microscope auxiliary components, such as electronic light shutters, filter wheels, motorized stages, and focus drift correction mechanisms, which can be coordinated and controlled by a host computer using commercially available image acquisition software. Electronic shutters are necessary to block illumination of the specimen between camera exposures when imaging fluorescently labeled cells in order to minimize photodamage, phototoxicity, and photobleaching, thus dramatically extending the quality of images and cell viability over the long periods of time often used in time-lapse experiments. Simultaneous imaging of multiple fluorophores, as well as combined fluorescence and DIC imaging, requires motorized filter wheels that can rapidly switch between several fluorescence filters and/or a polarizer. In practice, excitation illumination is controlled by a filter wheel that contains two or more bandpass excitation filters, while fluorescence emission is gathered through a second filter wheel that houses either bandpass or longpass barrier (emission) filters. Beneath the objective, a multiple bandpass dichromatic mirror is installed to coordinate blocking and reflecting of excitation wavelengths and simultaneous passing of emission wavelengths through a single, stationary optical element. Advanced filter wheels can switch between adjacent filters in 30 to 50 milliseconds with electronic shutters having operating specifications in the same range, which is the limiting factor in determining the shortest possible time interval between successive images in time-lapse sequences.
In confocal microscopy, time-lapse image collection is limited by the speed of the scanning mirrors. The highest scanning rates are achieved by reducing the image pixel dimensions and employing the fastest raster speed available on the instrument, but this is usually restricted to approximately 8 to 10 frames per second. Collecting image sequences over shorter time intervals requires single-line scanning, swept-field instruments, or spinning disk microscopes. Modern spinning disk and swept-field microscopes can routinely capture image sequences at high rates, ranging from one to several hundred frames per second, and are usually only limited by the speed of the digital camera system or photomultiplier. These instruments are designed to accommodate a host of experimental variables in high-speed time-lapse imaging at very low light levels.
Live-cell imaging experiments can be extraordinarily powerful, but they can also significantly benefit from complementary investigations using fixed cell assays to assist in the validation of phenomena that are observed in the living cells. Although this may seem counterintuitive due to the fact that live-cell assays are often held as the standard for cellular and molecular dynamics, the technical difficulties and risk of damaging cells during investigations can be substantially overcome if similar results in terms of kinetics and time point events can be verified with fixed cells. In many cases, comparing fixed cells to live-cell imaging is simply not possible, either because the kinetics of an event are too fast or because the nature of the experiment (for example, dynamics) cannot be meaningfully performed within the fixed cell. Moreover, the point of the live-cell experiment is to reveal events or properties that are not observed or easily interpreted in fixed cells. Nonetheless, it is worthwhile to consider the use of this approach as a technique for monitoring events seen in longer time-lapse experiments to confirm the absence of deleterious effects by extended illumination.
Modern instrumentation enables the imaging of living specimens at high signal-to-noise ratios using extremely low levels in incident illumination and now stands as a powerful technique for the analysis of molecular dynamics within cells. Continued advances in imaging techniques and fluorescent probe design enhance the power of this approach and ensure its future as an important tool in modern biology. Technological and conceptual advances in instrumentation are also likely to push spatial and temporal resolution to new limits, as well as perfecting the modes of fluorescence microscopy currently in use. Several of the recently introduced approaches, such as stimulated emission depletion and 4Pi microscopy, appear to be promising techniques for live-cell imaging, but their potential for widespread use in biological applications has yet to be established, and there are limitations on the thickness of acceptable specimens. In the final analysis, however, the technical care and expertise required to conduct a successful live-cell imaging experiment with currently available instrumentation is considerable, and even with likely advances there remain numerous obstacles.
Michael E. Dailey - Department of Biological Sciences and Neuroscience Program, 369 Biology Building, University of Iowa, Iowa City, Iowa, 52242.
Daniel C. Focht - Bioptechs Inc., 3560 Beck Road, Butler, Pennsylvania, 16002.
Alexey Khodjakov and Conly L. Rieder - Wadsworth Center, New York State Department of Health, Albany, New York, 12201, and Marine Biological Laboratory, Woods Hole, Massachusetts, 02543.
Kenneth R. Spring - Scientific Consultant, Lusby, Maryland, 20657.
Nathan S. Claxton, Scott G. Olenych, John D. Griffin, and Michael W. Davidson - National High Magnetic Field Laboratory, 1800 East Paul Dirac Dr., The Florida State University, Tallahassee, Florida, 32310.
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We characterized vaccinia IMV entry into HeLa cells and showed that the virus infection process has several interesting features. First, vaccinia IMV required actin polymerization prior to cell entry. That virus “surfing” requires normal actin dynamics has been reported for some retroviruses and vesicular stomatitis virus (36). Surfing may turn out to be a common transport mechanism shared by many viruses if vaccinia virus is added to the list of those using it. Plasma membrane ruffling and actin protrusions regulated by actin dynamics were also involved in vaccinia IMV recruitment. Next, we showed that IMV particles were endocytosed into HeLa cells in a manner independent of clathrin- and caveola-mediated pathways but dependent on dynamin, suggesting a virus entry pathway through fluid phase endocytosis (7). The role of endogenous dynamin 2 in vaccinia IMV entry into HeLa cells was demonstrated using three experimental approaches, first with an expressed DN-Dyn1 isoform, next with a dynamin-specific inhibitor, dynasore, and finally with a siRNA approach. In our hands, DN-Dyn2 blocked transferrin uptake but not IMV entry we suspect that a higher level of DN-Dyn2 is needed to block virus entry than to block ligand uptake. Alternatively, endogenous dynamin in HeLa cells may contain splicing variants that interacted with DN-Dyn1 and DN-Dyn2 with different affinities. It is worth noting that, despite a clear function of dynamin in clathrin- and caveola-mediated coated-vesicle formation, its role in coat-independent endocytosis processes such as fluid uptake has been controversial (1, 17, 18, 23, 24, 35). Four splicing variants of dynamin 2 have been reported (8, 54), and the DN GTPase mutants of all four splicing variants blocked transferrin uptake well however, only two splicing variants blocked dextran uptake, suggesting differential functions of the distinct splicing variants in fluid uptake (7). Furthermore, we identified and characterized VPEF, a novel cellular protein that is important for vaccinia IMV penetration into HeLa cells. Our data also suggested that VPEF and dynamin mediate IMV penetration through different mechanisms. How VPEF interacts with viral components to allow penetration remains to be clarified in the future. VPEF mRNA was detected in many mammalian cell lines (data not shown), suggesting a conserved function for VPEF. This would not be too surprising, given the wide host infectivity of vaccinia virus while we demonstrated the importance of VPEF in HeLa cells, it remains to be determined whether the same conclusion will apply to other cell types. It also awaits more experiments to determine whether the extracellular form of vaccinia virus requires VPEF for cell entry or not.
Because vaccinia virus attachment to cells at 4ଌ was relatively inefficient and only 1 to 2% of the input viral particles bound to cells after a 60-min incubation (37), we had to use relatively high-MOI (MOI = 40 to 100) infections in the EM and confocal analyses. Using viral early gene expression assays, we observed that vaccinia IMV entry into HeLa cells remained sensitive to BFLA in low-MOI (MOI = 0.1 to 5) infections, supporting the idea that MOI per se did not dictate the cell entry pathways employed by viruses. It is also worth noting that endocytosis of the WR strain of vaccinia IMV into HeLa cells, as described here, is not simply an artifact of high-MOI infections since infections of cells with the IHD-J strain of vaccinia IMV at a high MOI of 200 to 300 did not result in obvious endocytosis into BSC-1 (58) and HeLa cells (37).
In summary, these results demonstrate that vaccinia IMV is able to pirate cellular actin to get closer to cell bodies and manages to enter cells by hijacking a ubiquitous VPEF-dependent fluid transport pathway that is part of these cells' normal repertoire. We hypothesize that VPEF represents a new cargo or component of the earliest acceptor compartment of the clathrin- and caveolin-independent vesicle trafficking system (41) for the uptake of bulk phase fluid. The verification of this hypothesis will require further study.
Results and Discussion
MT-interaction differs significantly between P3 from pathogenic and non-pathogenic RABV
To assess the relationship of P3-MT-association with pathogenicity we expressed P3 from the Ni and Ni-CE RABV strains fused to green fluorescent protein (GFP Fig. 1 ) in a panel of mammalian cell lines from neuronal and extraneural tissues of several RABV susceptible/host species 23 ,24 ,25 ,26 , including COS-7 (African green monkey kidney), NSC-34 (mouse motor neuron-like), SK-N-SH (human neuronal), HeLa (human epithelial) and NA (mouse neuroblastoma). Expression of the proteins was confirmed by Western analysis lysates of COS-7 cells (see supplemental Fig. S1).
P3 is an N-terminally truncated isoform of P protein comprising residues 53, which is generated in infected cells by ribosomal leaky scanning that results in translation from an internal AUG codon corresponding to M53 (arrow) of the full length P protein (called P1) 10 . The CTD (containing the MT, N-RNA and STAT1 binding regions) and NTR (containing the dimerization domain) are indicated residue positions are indicated beneath the P1 protein. Residues at positions 56, 58, 66, 81 and 226 differ between P3 from the pathogenic (Ni) and attenuated (Ni-CE) strains of RABV (substitutions in Ni-CE-P3 are in red).
We previously found that MT interaction by GFP-fused P3 of the RABV CVSII strain can be detected in living cells by CLSM as association with a cytoplasmic filamentous network 19 . Consistent with this, extensive interaction of GFP-Ni-P3 with cytoplasmic filaments was observed in all cell types tested (COS-7, NSC-34, shown in Fig. 2a , and SK-N-SH, HeLa, NA, not shown). Importantly, GFP-Ni-CE-P3 filament association was clearly impaired in each of these cell types and comparable differences were observed between non-fused Ni-P3 and Ni-CE-P3 in fixed immunostained COS-7 cells (see supplementary Fig. S2). The Ni-P3-associated filaments were confirmed to be MTs based on sensitivity to MT-targeting drugs ( Fig. 2b ), and colocalization with mCherry-tubulin in living cells ( Fig. 2c ) and endogenous tubulin in immunostained cells ( Fig. 2d ). Thus, MT association differs between P3 proteins from pathogenic and attenuated virus.
(a) COS-7 cells were transfected to express the indicated proteins before analysis by live-cell CLSM each image is representative of cells in 30 fields of view sampled over 3 separate assays (COS-7) or 9 fields of view (NSC-34). (b𠄽) COS-7 cells transfected to express GFP-Ni-P3 were treated with or without Taxol or nocodazole (b) co-transfected to express mCherry-tubulin (c) or fixed and immunostained for β-tubulin (d) before analysis by CLSM colocalization in b and d is apparent as yellow coloration in merged image. (e) Live COS-7 cells expressing the indicated proteins were analyzed by CLSM to generate deconvoluted 3D images (images show reconstructed 3D images viewed down the z-axis). (f) Images such as those shown in (e) were analyzed to derive mean filament length values (mFL ± SEM n ≥ cells from 3 identical assays). p values were determined using the Mann Whitney test.
The Ni-CE P gene contains mutations compared with Ni P that result in 5 amino acid residue substitutions, one or more of which account for the differing function in IFN-antagonism and pathogenicity 13 ,20 ,21 . All of the substitutions affect the P3 sequence ( Fig. 1 ), resulting in four proline substitutions clustered within the N-terminal region (NTR), and one histidine substitution (N226-H) in the globular C-terminal domain (CTD). Previous mapping studies indicated that the latter domain mediates association with MTs 19 . To examine the effects of specific mutations, we expressed GFP-Ni-P3 containing N226-H alone (GFP-Ni-P3-N226-H) or the NTR mutations (GFP-Ni-P3-CENTR) for analysis as above. MT filament association of GFP-Ni-P3-N226-H, but not GFP-Ni-P3-CENTR, was clearly impaired in COS-7, NSC-34 ( Fig. 2a ), and SK-N-SH, HeLa, and NA cells (not shown).
To quantify the extent of MT network association, and effects thereon, of Ni-CE mutations we generated deconvoluted 3D images of living cells expressing GFP-fused P3 proteins (images viewed down the z-axis are shown in Fig. 2e ), and developed a methodology to detect and measure intracellular filamentous GFP using the Imaris software filament-tracing algorithm tool (see Materials and Methods) this is to our knowledge the first application of this tool to quantify protein-MT interaction in living cells. This enabled semi-automated detection of P3-associated filaments in single cells to calculate the total filament length. The corresponding mean filament length (mFL) per cell was then determined for ≥ 30 cells for each protein tested ( Fig. 2f ). Results confirmed that MT-association of GFP-Ni-CE-P3 and GFP-Ni-P3-N226-H, but not GFP-Ni-P3-CENTR, was significantly reduced compared to GFP-Ni-P3 (p <𠂐.0001 and p =𠂐.0118 for Ni-CE and Ni-P3-N226-H, respectively), with the N226-H mutation alone causing a c. 50% decrease in the mFL of Ni-P3. Thus, N226-H, but not the NTR mutations, directly impacts on P3-MT association. However, since N226-H alone does not fully recapitulate the phenotype of Ni-CE-P3 ( Fig. 2f ) it appears the NTR mutations can augment the effect of N226-H, indicative of an indirect/regulatory role of the NTR.
To confirm that the reduced mFL for Ni-CE-P3 and Ni-P3-N226-H is due to decreased interaction with MTs, we performed cytoplasmic extraction of transfected cells to remove non-MT associated protein, as described 27 , before immunostaining for MTs and CLSM analysis. Quantitation of P3-MT colocalization using Pearson’s coefficient indicated that MT-association is significantly (p <𠂐.0001) impaired for GFP-Ni-CE-P3 and GFP-Ni-P3-N226-H compared with GFP-Ni-P3 ( Fig. 3 ). However, the capacity of Ni-P3-N226-H to associate with MTs remained greater than that of Ni-CE-P3, consistent with data from the live-cell filament tracing assays ( Fig. 2f ).
N226-H mutation inhibits colocalization of P3 with MTs (a) COS-7 cells expressing the indicated proteins were extracted to remove soluble (non-MT-associated) protein before fixation and immunostaining for β-tubulin, and analysis by CLSM (colocalization is apparent as white coloration in merged image). (b) Images such as those shown in (a) were analyzed to derive Pearson’s coefficient as a measure of colocalization between GFP-P3 and β-tubulin (mean ± SEM n ≥ 14 cells from two assays). p values were determined using Student’s t-test or the Mann-Whitney test.
MT bundling by P3 is significantly impaired by N226-H mutation
Interactions of cellular and viral MAPs with MTs often induce gross structural changes in MT networks that can be detected by light microscopy/CLSM analysis and are thought to correspond to MT bundling 28 ,29 ,30 ,31 ,32 ,33 ,34 ,35 . Such effects could provide a means to quantify MAP-MT interactions at the level of individual MT filaments. However, light microscopy/CLSM is diffraction limited with the greatest possible spatial resolution c. 200 nm, precluding direct quantitative analysis of individual MT filaments (c. 25 nm diameter) or of the extent of filament bundling. While electron microscopy (EM) can visualize individual filaments, and has been used to confirm bundling activity of certain MAPs 28 ,29 ,30 ,31 ,32 ,33 ,34 ,35 , such studies are typically limited to complexes of purified protein with in vitro polymerized tubulin, or analysis of cells subjected to extensive chemical fixation procedures/sample preparation times 36 ,37 . Furthermore, EM precludes the use of fluorescent tags for precise localization of structures/proteins of interest in large cell populations. Thus, to directly detect and quantify cellular MT filament bundling induced by P3, we utilized dSTORM super-resolution imaging, which overcomes the diffraction limit by detecting unperturbed emission point spread functions from single molecules, enabling highly precise spatial localization of the emitting fluorophores. Analysis used cell fixation and immunostaining protocols known to preserve MT integrity, which we have been used previously to visualize individual MTs 38 ,39 ,40 , with dSTORM measurements made using a custom-built super-resolution widefield microscope that can achieve single molecule localization precisions better than 10 nm, and spatial resolution down to 20 nm 41 . Detailed images of MT architecture were acquired with sufficient resolution to differentiate closely associated MTs (presumed to be bundles, see below) from individual MTs which are tens of nanometers apart but occupy the same diffraction limited area and so cannot be resolved using conventional light microscopy (see supplementary Fig. S3).
To quantitatively analyze the dimensions of the filaments and occurrence of bundling in dSTORM images, we calculated the relative MT feature diameter (MTfd) using the width at half-height of a Gaussian function fit to the average intensity profile of a cross-section of a continuous MT feature ( Fig. 4a ). The width of single MT filaments imaged by this method will be greater than the intrinsic diameter of a MT filament (c. 25 nm) since it is the locations of antibody-conjugated Alexa Fluor-647 molecules that are detected. Thus, the primary immunolabel against tubulin and the Alex Fluor 647-conjugated secondary immunolabel (which label both sides of the filament) add some 30 nm to the apparent width of a single MT. Taking this and the localization precision of the fluorophores themselves (typically 9 nm in these measurements) into account, we predicted a range of 40 nm for single MTs. In agreement with this, benchmarking using mock-transfected cells indicated that 72% of MTfds were in this range (see supplementary Fig. S3). The thicker filaments detected in mock transfected cells ( nm) are likely the result of native bundling of MTs as well as detection of overlapping MTs that are separated in axial space but appear to associate in the 2D dSTORM image. Nevertheless, the majority of features are within the range expected for a population of largely individual, non-bundled MTs.
(a) dSTORM images of immunostained β-tubulin in COS-7 cells expressing the indicated proteins are shown in the upper panels. Boxed areas are expanded in the lower panels, and Gaussian functions (black line) fit to the average intensity profile (red line) of the indicated filaments shown below. The derived MTfd (calculated at the full width half-height of the Gaussian function) is indicated. (b) The frequency distribution of MTfds calculated for each protein is shown (n = [GFP-Ni-P3], 1071 [GFP-Ni-CE-P3] and 1299 [GFP-Ni-P3-N226-H] measurements are from 10 cells for each protein over two identical assays). (c) Scatter dot plots of MTfds shown in (b). p values were determined using the Mann-Whitney test.
Analysis of GFP-Ni-P3-expressing cells ( Fig. 4a ) indicated a substantial change in MT architecture, with 63.4% of MTfds exceeding 40 nm, and 18.1% exceeding 200 nm ( Fig. 4b ), indicative of extensive bundling. The notion that these large MT features correspond to multiplex bundles of single MT filaments is supported by the observation that the thicker filaments often appeared to split into thinner filaments, with the continuous length of the 𠇋undled” region indicative of specific association of the constituent MTs (see supplementary Fig. S4).
The architecture of MT networks in cells expressing GFP-Ni-CE-P3 and GFP-Ni-P3-N226-H ( Fig. 4a ) was much more consistent with that observed in mock-transfected cells than in cells expressing GFP-Ni-P3, with 78.5% (GFP-Ni-CE-P3) and 67.3% (GFP-Ni-P3-N226-H) of MTfds within 40 nm ( Fig. 4b ), indicative of defective bundling by these proteins. However, Ni-P3-N226-H retained a slightly increased bundling capacity compared to Ni-CE-P3, indicated by a >𠂒 fold higher proportion of MTfds in all bins of the frequency distribution exceeding 140 nm. Consistent with the observed trend, statistical analysis indicated that the median MTfd for cells expressing GFP-Ni-P3 was significantly (p <𠂐.0001) greater than that for GFP-Ni-P3-N226-H, which in turn was significantly (p <𠂐.0001) greater than that for GFP-Ni-CE-P3 ( Fig. 4c ).
Thus, it appears that N226-H substantially inhibits bundling, but does not fully recapitulate the defective phenotype of Ni-CE-P3, so that the degree of bundling detected by dSTORM correlates with the extent of MT network association determined in CLSM-based filament tracing and co-localization assays ( Figs 2 and and3). 3 ). These data indicate that dSTORM provides an effective method to quantify protein-MT association at the level of individual filaments. This is to our knowledge the first application of dSTORM to interrogate viral protein-MT interaction and highlights the power of dSTORM to elucidate fundamental processes at the virus-host interface. Together, the data indicate that N226-H directly impacts on P3-MT-association, consistent with its localization to the CTD that contains MT-association activity ( Fig. 1 ) 19 .
Interaction of P3 with RABV N-protein and homodimerisation of P protein is not impaired by mutations in Ni-CE
P protein has critical roles in viral genome replication, which requires interaction of the CTD with RABV nucleoprotein (N protein) to mediate association of P protein with N-protein-encapsidated genomic RNA (N-RNA Fig. 1 ) 42 . The CTD also mediates interaction with STAT1 ( Fig. 1 ), which is important to the IFN-antagonistic functions of P protein isoforms 15 ,16 ,43 ,44 . The reported N- and STAT1-binding sites are distinct from N226 44 ,45 ,46 , and previous data indicate that they are not significantly impacted by N226-H, as Ni-CE virus is highly viable in cultured cells, and Ni-CE-P is not deficient for STAT1 interaction 13 . To confirm directly that Ni-CE-P CTD interaction with N-protein is unimpaired, we used yeast-2-hybrid analysis ( Fig. 5 ), which has been used extensively to identify and characterize bi-molecular interactions of P protein, including with N protein, as well as P protein homodimerisation 47 ,48 ,49 . This indicated no evident defect for Ni-CE compared with Ni P proteins. 2-hybrid analysis of homodimerization of P protein via the NTR-localized dimerization domain 50 ( Fig. 1 ), which has been shown to be required for P3-MT association via the CTD and for MT-dependent inhibition of STAT1 19 , also indicated no defect in Ni-CE-P ( Fig. 5 ). Thus, the mutations in Ni-CE do not appear to be generally detrimental to P protein structure or function, indicative of specific effects on MT interaction such that the N226-H mutation could be applied to investigate directly the functional significance of P3-MT association.
L40 yeast cells were cotransformed to express the DNA-binding domain of LexA (Lex) or the GAL4-activation domain (GAD) fused to Ni-P1, Ni-CE-P1, Ni-P3, Ni-CE P3, the corresponding P3 NTRs (P54-172), or RABV N protein. Growth streaks are shown, where interaction of Lex- and GAD-fused proteins is indicated by dark coloration of colonies.
N226-H mutation impairs IFN antagonist function of P3 and pathogenicity of RABV
To examine the effect of N226-H mutation on the capacity of P3 to cause association of STAT1 with MTs, cells expressing GFP-fused Ni-P3, Ni-CE-P3 and Ni-P3-N226-H were treated with IFNα before cytoplasmic extraction, fixation and immunostaining for STAT1 and tubulin, and imaging by CLSM. Association of GFP-Ni-P3 and STAT1 with MTs was clearly detectable ( Fig. 6a ), but association of STAT1 with MTs was strongly reduced or undetectable in cells expressing GFP-Ni-P3-N226-H, and there was no evidence of STAT1-MT interaction in cells expressing GFP-Ni-CE-P3 or control cells transfected to express GFP alone. To further analyze the effect of N226-H on P3-mediated antagonism of IFNα/STAT1-dependent signaling, we used a luciferase reporter gene assay, as previously described 13 ,19 ,44 . Ni-P3 strongly inhibited signaling in response to IFNα, but Ni-CE-P3 and Ni-P3-N226-H were significantly (p <𠂐.0001) impaired in this respect ( Fig. 6b ). Notably, the IFN-antagonistic function of the different P3 proteins correlated with their capacity to associate with MTs and induce MT-association of STAT1 ( Figs 2 , ,3, 3 , ,4 4 and and6a), 6a ), and with the pathogenicity of Ni and Ni-CE viruses.
(a) IFNα-treated COS-7 cells expressing the indicated proteins were extracted before fixation and immunostaining for STAT1 and β-tubulin, and analysis by CLSM images are representative of 15 fields of view from two assays. (b) IFN-α-dependent signaling in COS-7 cells expressing the indicated proteins was analyzed using a dual luciferase reporter gene assay, as previously described 13 ,19 ,43 . Luciferase activity is expressed as fold change relative to that obtained for IFN-α-treated cells expressing Ni-P3 protein (mean relative light units [RLU] ± SEM n = from 4 identical assays). p values were calculated using Student’s t-test.
To directly investigate the effect of N226-H on infection in vivo, we introduced this mutation alone into the Ni P gene of the pathogenic IFN-resistant CE(NiP) virus 13 ,21 the derived recombinant virus was called CE(NiP-N226-H) ( Fig. 7a ). ddY mice (five mice per virus) were inoculated intracerebrally with 100 focus forming units (FFU) of Ni-CE, CE(NiP) or CE(NiP-N226-H) virus, and monitored over 21 days post-infection (dpi), as previously 13 . Consistent with previous observations, mock infection was asymptomatic (data not shown), with infection by Ni-CE virus causing only mild and temporary weight changes, while CE(NiP) caused death or sacrifice at the defined end-point in 1 out of 5 mice by 11 dpi, and 5/5 mice by 14 dpi ( Fig. 7b ). The outcome of infection with CE(NiP-N226-H) was similar to infection with Ni-CE, causing only minor temporary weight loss with no onset of major symptoms and no fatalities, indicating significant attenuation (p =𠂐.0027). We also performed infection using 10 6 FFU virus, finding that infection by Ni-CE remained non-lethal while all CE(NiP) infected mice succumbed to infection by 8 dpi. Significant (p =𠂐.0052) attenuation of CE(NiP-N226-H) was observed an the increased FFU, with only 3/5 mice succumbing by 14 dpi while the remaining mice showed increasing weight from 11 dpi.
(a) Schematic representation of the genomes of viruses used genes from Ni and Ni-CE are in black and white, respectively. The P gene of Ni-CE is substituted for the P gene of Ni in CE(NiP), and for a mutated version of the Ni P gene containing N226-H in CE(NiP-N226-H). (b,c) 100 (b) or 10 6 (c) FFU of the indicated virus was inoculated intracerebrally into mice (five mice per condition) and body weight relative to that at 0 dpi (mean relative body weight ± SEM, for live mice) and survival was monitored over 21 or 14 dpi. p-values for survival curves were calculated using log-rank (Mantel-Cox) test. † all mice dead or sacrificed on reaching end point.
Taken together, our data indicate that N226-H mutation impairs the capacity of P3 to interact with MTs and effect antagonism of antiviral signaling, and strongly reduces the ability of RABV to cause lethal infection in mice. These findings are consistent with key roles for MT-dependent IFN-antagonism through the P3-MT interface in the pathogenicity of RABV, supporting the hypothesis that viral protein-MT interactions have significant roles in subversion of host biology distinct from well-established functions of MTs in virus trafficking 9 . Importantly, the observation that modification of these interactions can affect disease outcomes in vivo identifies new potential targets at the virus-MT interface that could contribute to the development of attenuated vaccines and/or antiviral drugs to combat infection by lyssaviruses, which cause rabies disease with an almost 100% case-fatality rate and no effective therapeutic options 51 .
Advanced Image Acquisition and Analytical Techniques for Studies of Living Cells and Tissue Sections
Studies on fixed samples or genome-wide analyses of nuclear processes are useful for generating snapshots of a cell population at a particular time point. However, these experimental approaches do not provide information at the single-cell level. Genome-wide studies cannot assess variability between individual cells that are cultured in vitro or originate from different pathological stages. Immunohistochemistry and immunofluorescence are fundamental experimental approaches in clinical laboratories and are also widely used in basic research. However, the fixation procedure may generate artifacts and prevents monitoring of the dynamics of nuclear processes. Therefore, live-cell imaging is critical for studying the kinetics of basic nuclear events, such as DNA replication, transcription, splicing, and DNA repair. This review is focused on the advanced microscopy analyses of the cells, with a particular focus on live cells. We note some methodological innovations and new options for microscope systems that can also be used to study tissue sections. Cornerstone methods for the biophysical research of living cells, such as fluorescence recovery after photobleaching and fluorescence resonance energy transfer, are also discussed, as are studies on the effects of radiation at the individual cellular level.
Plant cell wall-derived biomass serves as a renewable source of energy and materials with increasing importance. The cell walls are biomacromolecular assemblies defined by a fine arrangement of different classes of polysaccharides, proteoglycans, and aromatic polymers and are one of the most complex structures in Nature. One of the most challenging tasks of cell biology and biomass biotechnology research is to image the structure and organization of this complex matrix, as well as to visualize the compartmentalized, multiplayer biosynthetic machineries that build the elaborate cell wall architecture. Better knowledge of the plant cells, cell walls, and whole tissue is essential for bioengineering efforts and for designing efficient strategies of industrial deconstruction of the cell wall-derived biomass and its saccharification. Cell wall-directed molecular probes and analysis by light microscopy, which is capable of imaging with a high level of specificity, little sample processing, and often in real time, are important tools to understand cell wall assemblies. This review provides a comprehensive overview about the possibilities for fluorescence label-based imaging techniques and a variety of probing methods, discussing both well-established and emerging tools. Examples of applications of these tools are provided. We also list and discuss the advantages and limitations of the methods. Specifically, we elaborate on what are the most important considerations when applying a particular technique for plants, the potential for future development, and how the plant cell wall field might be inspired by advances in the biomedical and general cell biology fields.
Plant growth is based on the ability of plants to convert carbon dioxide into sugars via photosynthesis and metabolize them into a wide range of other biomolecules . The main carbon sink in plants is the cell wall an extracellular matrix composed of long-chain glycans, glycoproteins, phenolic and polyester polymers, as well as solutes and water. The cell walls provide important structural and protective functions to plants as well as contribute the bulk of their biomass . A large number of products, such as biobased fuels, chemicals, paper, and novel materials may be derived from this biomass, and finding sustainable and carbon-neutral approaches to do this will be an important part of shifting our society away from a fossil-fuel based economy . We anticipate that these efforts will be aided by a better understanding of how this biomass is structured, how it is created by the plant, and what happens as it is being processed, and discuss the toolset available to accomplish these studies using fluorescence microscopy in this review.
The major constituents of plant cell walls are polysaccharides, which are divided into three different classes: cellulose, hemicelluloses, and pectins [4𠄶]. Cellulose consists of β -1,4-linked glucose that coalesces into microfibrils via intermolecular hydrogen bonds and van der Waal’s forces. The cellulose microfibrils have a high tensile strength and work as a scaffold, providing the load-bearing strength to the cell walls [7, 8]. Cellulose is produced at the cell surface by cellulose synthase (CesA) protein complexes (CSCs), which utilize cytosolic UDP-glucose as their substrate [9, 10].
Hemicelluloses primarily consist of β -1,4-linked neutral sugar backbones with equatorial conformations and include xyloglucan, xylan, mannan, glucomannan, and mixed-linkage glucan . These polymers are made in the Golgi lumen, with the possible exception of mixed-linked glucan , by glycosyltransferases (GTs) that use an array of nucleotide sugars as substrates. Hemicelluloses engage with cellulose and/or lignin to regulate, depending on the developmental context, either cell wall expansion and cell growth or cell wall rigidification [8, 14, 15].
Pectins are also made in the Golgi lumen by GTs and are some of the most complex and dynamic cell wall molecules. Homogalacturonan (HG), a homopolymer of α -1,4-linked galacturonic acid, is synthesized in a highly methylesterified form and upon secretion in the apoplastic moiety can be de-esterified by a class of enzymes called pectin methylesterases (PMEs). The modulation of PME activity underlies cell wall-directed cellular and developmental processes, for instance, meristem formation or pavement cell morphogenesis . HG backbone can be decorated with monosaccharides such as apiose (apiogalacturonan), xylose (xylogalacturonan), or by a complex assortment of sugars and glycosidic linkages known as rhamnoglacturonan II (RG-II). Another pectin with a backbone of repeating disaccharide of galacturonic acid and rhamnose units is rhamnogalacturonan I (RG-I), which is further modified with galactan and arabinan side chains. Methylation and acetylation of pectins provide further important molecular features that influence biomass processing and fermentability [19, 20].
Unlike the flexible primary cell walls which encase cells that are still growing, thick secondary cell walls are deposited once cells have ceased growth. These strong walls provide mechanical strength as well as creating the vascular tissue needed for water transport and providing resistance to biotic threats . The secondary cell walls make up the bulk of a plant’s biomass and are the major source of fermentable sugars for cellulosic biofuel production . A prominent component of many secondary cell walls is lignin, which is a highly heterogenous phenolic polymer that is polymerized directly in the cell wall by laccases and peroxidase-assisted radical coupling of small aromatic alcohols known as monolignols . This extensive crosslinking reinforces the cell walls, but lignin itself also acts as an essential hydrophobic barrier on xylem vessels to enable long distance water transport.
Because of its abundance and extensive crosslinking, lignin is usually the main factor that influences the resistance of cell walls to decomposition. However, the recalcitrance of biomass to processing is still a poorly understood phenomenon that is also influenced by cell wall morphology, porosity, and the abundance of the varying constituent polymers . Fluorescence-based imaging can be used to assess these features in both native cell walls and processed samples, showing things like cell wall microdomains, the accessibility of enzymatic machineries, or the effects of physical or chemical treatments on the sample [28, 29].
There are a handful of imaging methods that utilize the intrinsic chemical or mechanical features of cell wall polymers, such as atomic force microscopy (AFM), Fourier-transformed infrared (FTIR) microspectroscopy, confocal Raman microspectroscopy (CRM), coherent Anti-Stokes Raman scattering microscopy (CARS), stimulated Raman scattering microscopy (SRS), Brillouin microscopy, and X-ray computed tomography (CT) [29, 30]. Although these techniques are outside of the scope of this review, they are expected to be particularly useful for characterization of samples with unusual functional groups or electronic densities. Additionally, some fluorescence-based methods like Förster resonance energy transfer (FRET) microscopy and fluorescence lifetime imaging microscopy (FLIM) have recently been well summarized in other reviews [30, 31], so we do not discuss the special biophysical information they can provide here but note that they share many of the same challenges as standard fluorescent imaging and molecular tagging.
This review is divided into two major sections: the first part discusses options for cell wall visualization by various types of fluorescence microscopy and the second section deals with molecules that enable exogenous or endogenous ‘tagging’ specific molecular targets. All research methods come with their own limitations, drawbacks, and challenges, and we use this opportunity to particularly highlight these. We also suggest how future advances and development in the field can mitigate the current drawbacks and pitfalls.
Figure 6. Confocal fluorescence images (75 μm × 75 μm) of Hoechst 33258 (nuclear stain) + 10 μM 1 after 24 h of incubation.
To summarize, the first example of a M–M bonded compound incorporating an organic fluorophore has been synthesized. The present results with compound 1 indicate that dirhodium compounds can be tagged with fluorescent probes and that the intracellular localization is dictated at least in part by the tethered metal complex since the cellular distribution pattern of 1 differs from that of the free phenbodipy ligand. Compound 1 was found to target mainly lysosomes and mitochondria at concentrations in the 1–100 μM range, with a slight preference for the former organelle (∼1.4-fold). In contrast to the closely related compound 2 (see molecular structure in Figure S1), which targets the nucleus and induces DNA damage, compound 1 does not localize in the nuclei of A549 cells, evidence that supports the contention that various cellular organelles can be targeted by tuning the ligands of the dirhodium unit. In this vein, further studies are underway in our laboratories to modify the nature and lipophilicity of the fluorophore, to change its position relative to the dirhodium core (equatorial binding versus covalently attached to the bridging carboxylate ligands) in order to improve the uptake and cytotoxicity of this new type of fluorescent dirhodium compound. Ultimately, the aim is to gain deeper insight into the anticancer properties of this interesting class of M–M bonded compounds. Moreover, the current study provides an impetus for probing the biological properties of other multicenter inorganic complexes, since the same strategy can be used to label diruthenium(28) and dirhenium(29) anticancer compounds. It is worth pointing out that the realization that Rh–Rh bonded compounds can be successfully tagged with light-harvesting units such as bodipy will positively impact other research areas, such as the use of dirhodium compounds as photocatalysts,(30-32) since attaching a moiety with a high molar absorptivity to the dimetal core is expected to improve the efficiency of such catalytic systems.
NIH Image : Use In Fluorescence and Confocal Microscopy
The work that led to this manual was supported by grants to H. J. Karten from NIMH, NINDS and NEI. This manual was prepared with the direct support of NIMH and the Human Brain Project and the Brain Database Project. Under the stress of attempts to obtain grant support, we often neglect to sufficiently thank the many dedicated scientists who work so hard to establish those programs that provide the necessary funds. This manual is dedicated to those many loyal supporters of all of us at NIH, NSF, DOD, NASA, and elsewhere.
I have adopted and adapted a number of macros contributed by various NIH Image users. The authorship of the original macros was often difficult to determine. I am grateful to the unacknowledged authors, and hope that they are not offended if not specifically credited. My special thanks to Rusty Gage and Larry Goldstein for providing me with unlimited access to their confocal microscope facilities. Without their generosity, this manual would not have been possible.
Please let me know if this manual is helpful. Please inform me of any of errors in this manual. If you have any suggestions to improve this manual, additional macros that are helpful for processing confocal images, or other strategies for processing CLSM images, please let me know, and I will try to incorporate these changes in future versions.
Harvey J. Karten, M.D.
NIH Image is a useful tool for evaluating fluorescent material prior to examining it on the confocal microscope and for post-processing of confocal laser scanning microscope (CLSM) images. The experienced user of NIH Image may find many of these operations obvious. However, many users of CLSMs, previously unfamiliar with NIH Image will find it a useful tool for both pre- and post-acquisition processing of images. I hope that this manual provides a relatively easy introduction to the use of NIH Image for CLSM users.
Performing post-acquisition processing of images on the Macintosh will free the confocal instrument for image collection. Though not as extensive as VoxelView, Analyze, VolVis, SYNU or VoxBlast, particularly in applications requiring advanced volume rendering and Voxel based calculations, the use of NIH Image and Adobe Photoshop 3.0 (Adobe Systems Inc., Mountain View, CA) on the PowerPC will provide most of the functions needed for manipulating confocal images, and at a much lower price.
This manual is intended for use with NIH Image , version 1.60 or later. Several of the procedures described in the manual will not operate correctly on earlier versions. You can obtain a copy of the latest version of NIH Image from the FTP site: zippy.nimh.nih.gov (login: anonymous Password: <your EMail address> cd/pub/image) or the associated Website (http://rsb.info.nih.gov/nih-image/).
1. NIH Image on Macintosh and PC/Windows 95
This manual describes the use of NIH Image on a Macintosh. In principle, it should be fully applicable to a version of NIH Image being prepared for PC/Windows 95.
Tod Weinberg of Scion Corporation (Frederick, MD) is sponsoring the transfer of NIH Image to PC/Windows95. This is a direct "port" of NIH Image --i.e., it should have the exact look and feel of NIH Image for the Macintosh, with the exception of those features of the operating system that reflect the individual platforms. At this time (October, 1996), ImagePC ( NIH Image for PC) is still in alpha testing, and some operations are not yet functional, but it is available for downloading by interested individuals. It is available from the same location that provides NIH Image for the Macintosh, both from the FTP site and the Website. The "port" of the program to the PC seems very promising, and the final version will hopefully permit users to perform the same operations on the PC and Macintosh with equal facility.
2. Contents Of This Manual
A) How to use NIH Image to evaluate the quality of single or multiple labeled fluorescent histological sections in preparation for confocal microscopy.
B) How to transfer files from the BioRAD, Leica or Zeiss CLSM to NIH Image , both individual images and Z-series.
C) How to use NIH Image for image processing and analysis: NIH Image provides a wide range of functions for image analysis and processing, including changing contrast and brightness values, pseudocoloring images, rotating, cropping, scaling, various filtering operations, measuring density, density slicing, measuring length and area, profile of a line, and counting particles. These are useful tools for modifying and analyzing confocal images. These functions fall into the more general category of image processing, and are discussed at greater length in the NIH Image manual, About NIH Image.
D) How to use multiple windows, Z-series and Stacks: The Macintosh allows you to open many windows, each containing a different confocal image. This facilitates comparison of different images. The various Stacks functions are amongst the most powerful features of NIH Image . NIH Image Stacks function allows you to rapidly step back-and-forth through a Z-series of sections. You can crop this Stack to select specific features of interest. You can also use the Stacks function to rapidly and alternately compare different images.
E) How to "Project" all the plates (or only a selected number of plates) of a Z-series onto a single plane.
F) How to generate 3D images and 3D rotations: NIH Image allows you to generate a 3D-series of images from the Z-series. You can display them in a montage window, fabricate stereo pairs or animate them to give the appearance of rotating a 3D object in space.
G) How to merge double and triple labeled sections, producing a three color images (RGB) with NIH Image .
H) How NIH Image can generate a double/triple labeled 3D-series for stereo pairs and rotations.
3. Some Hardware Considerations
Macintosh computers have proven particularly suitable for graphics applications. Their 32-bit memory model permits use of large quantities of RAM for rapid processing of image files. The graphics hardware and operating system have proven very suitable for image processing. All the procedures described in this manual can be achieved on a "low end" Macintosh, such as the Mac IIci, IIfx, to midlevel machines such as the 68040 Quadra series and, of course, will perform extremely well on the PowerPC series of machines. Most of the routines described in this manual were developed using a Quadra 950. Separate versions of NIH Image , optimized for the 680X0 series, as well as a single "fat" binary version that is optimized for both a 680X0 and a PowerPC, are available. Both versions are now available from the FTP site. Much less expensive than many dedicated graphics workstations, such as a Silicon Graphics Indy^2, the cost of a fully equipped Macintosh will vary in price based on some of the variables listed below. NIH Image will perform adequately on any current Macintosh.
Major differences in performance will be noticed when generating a rotation series, resectioning a Z-series, or applying various filters. It is not necessary to have a 68040 with a floating point processor (FPU). However, for improved speed performance, a PowerPC 7100, 8500, or the 9500, will prove most attractive. The 6100 does not have room for a frame grabber or a video output card to video printers or VHS tape. If you have the funds, the 9500/200 MHz unit is obviously preferable. The 8500 and 9500 have fast SCSI-2 ports for rapid data file transfers between external devices, built-in Ethernet for communication, and can be equipped with a high speed graphics display card. NIH Image does not support a multiprocessor Macintosh.
As we accumulate more experience using NIH Image on a PC/Windows 95 platform, future editions of this manual will include comments pertinent to that platform. Based on preliminary considerations, we suggest the use of a fast Pentium (e.g., Pentium Pro 200), with large quantities of RAM, fast 24-bit graphics card, a large hard disk and a 20-inch RGB monitor.
A) Operating System
Use System 7.1 or higher. System 7.5.5 is quite stable, and we now use it routinely.
B) Graphic RAM and Monitors
NIH Image is an 8-bit program. However, beginning with version 1.56 NIH Image will run even if the monitor is in 24-bit mode. This permits rapid switching between Adobe Photoshop 3.0 (24-bit program) and NIH Image . In order to obtain a full 24-bit image on a 20-inch monitor (at 1024x768 or higher), the graphics display card on an PowerPC should be packed with 4 MB of VRAM to permit 24-bit color on a 20-inch monitor. The Apple Trinitron 20-inch multisynch monitor provides an excellent image, and also is the lowest priced 20-inch monitor for the size and quality of the image. However, all the operations outlined in this manual will work on a 14-inch monitor with 8-bits. The 7100 and 6100 only permit 2 MB of VRAM, thus you will be limited to using a 17-inch monitor with 24-bit color.
C) RAM Vs. Virtual Memory
You will need large quantities of RAM for optimal performance. I suggest a minimum of 32 MB of RAM, and more if you can afford it. Avoid Virtual Memory if at all possible. (However, Apple does recommend that you assign 1 MB of Virtual Memory when using System 7.5.x on a PowerPC.) Graphics files are large, and confocal Z-series are often huge. If you also have a motorized stage and make extended XY planes of Z-series, your files are likely to be 30-50 MB or larger and you will have to reconcile yourself to buying a minimum of 80-256 MB of RAM if you want to move faster than a glacier. A minimal rule of thumb is that you should have 2.5-3 times more RAM than the size of your largest files. You should also learn how to allocate this memory to NIH Image using Get Info . Otherwise the additional memory will be of no benefit to you. I suggest that you allocate a minimum of 24 MB of RAM to NIH Image , and another 24 MB to Adobe Photoshop. Play around with the program. Many people are using RAM Doubler to compensate for limited amounts of RAM. I have no experience with this program and have heard mixed reports as to it value in working with large graphics files.
D) Disks And Storage Media
You will need storage media with a large capacity. But I assume that if you are working with confocal images, you have already had to deal with this problem. A hard disk of at least 500 MB is required. For greater flexibility, a system disk of 1 GB and a magneto- optical disk of 500 MB or larger for your data files is suggested. You may find it helpful to transfer your larger files from the slower magneto-optical to your system hard disk when working on them. Then move them back to the magneto-optical for long term storage. Confocal Z-series may readily exceed 25 MB. You should have enough storage space on your hard disk to allow you to save both the original file and any modifications you may make during a work session. We often find that we use more than 75 MB per Z-series in a single session.
A few words about those clever System Extensions on the Macintosh--the Extensions that make tea while you compute, or respond to voice commands (such as Apple's PlainTalk). These extensions not only chew up space, but worse, they convert your PowerPC to molasses while continually polling for voice inputs, losing CPU cycles, etc. Turn them all off. Stay with the basics. When you are doing image processing, turn off "File Sharing". If someone else on the network decides to just glance at your directory, your machine will be distracted and slow things down. You will still be able to get back onto the network with the click of key, but not be impeded by curious onlookers. Get used to dedicated computing, as in the days of yore, if you want to obtain your results as speedily as possible.
4. Loading NIH Image And The Confocal Macros
This manual is written with the assumption that the reader is familiar with the Macintosh Operating System 7.1 or higher.
Make sure that you have allocated sufficient memory to NIH Image to perform many of the operations outlined. In addition to allocating memory to the program as described above, using the File menu item, Get Info, you must also allocate sufficient memory to the NIH Image "Undo & Clipboard Buffer Size", shown under the Options + Preferences menu item. I suggest a minimum of 1400 KB. You must then Record Preferences under the File menu. Close the program and Restart the program.
The accompanying file, "Confocal Macros" should be copied into the Folder (Directory) containing the other NIH Image macros. Select the Specials menu, and the Load Macros item. Select the "Confocal Macros" from the resulting dialog box.
The "Confocal Macros" file contains a series of macros that are particularly useful for processing double labeled pairs of sections, Z-series, and generating stereo pairs. The reader should also familiarize themselves with the use of Stacks in NIH Image , as described in About NIH Image.
A copy of the macros file is included in the binhexed file "/pub/image/documents/confocals.hqx" provided via FTP. However, for those users who have obtained this file directly from the Website, the confocal macros file is not readily available. In order to provide access to this file, "Confocal Macros" has been appended to the end of this manual. It should be copied to a "Simple Text" file, and saved as "Confocal Macros."
5. Other Software Used In Conjunction With NIH Image
NIH Image provides a variety of powerful operations not readily available in other software, regardless of the price. Some functions, however, are best performed using other software packages. This is particularly true for those procedures that benefit from true 24-bit operations, such as adjusting individual color planes, color filtration and printing 24-bit images to dye sublimation printers.
A) Adobe Photoshop 3.0.5 and Canvas 5.0
Most confocal microscopy labs currently use Adobe Photoshop 3.0 for adjusting color saturation, brightness/contrast, cropping, assembling composite illustrations of multiple images, labeling the images and printing RGB confocal images. A new version of Canvas 5.0 (Deneba Corporation, Miami, FL) provides all these important functions of Adobe Photoshop needed by confocal microscopists for about one-third of the cost. Canvas 5.0 handles text labels and layers much better than Adobe Photoshop, but the cropping tool takes a bit of getting used to.
B) FileMaker Pro 3.0
In order to maintain a record of the content of each image or series of images as you collect them on the confocal scope, I have prepared a FileMaker Pro template. The Template is also posted on the FTP server for NIH Image within "confocals.hqx". When you download and decompress "confocals.hqx" the template will be decompressed as "Confocal_FileMaker_Template". FileMaker Pro is a simple and inexpensive database program. It is available for both Macintosh and DOS/Windows, and files are interchangeable between the two versions.
The careful design of a database will encourage you to store important information including the material and parameters used for the collection of the original image, storage location of the file, and modifications to the file.
6. Image Database Software
FileMaker Pro 3.0 can save images as well as text. However, dedicated image databases will automatically scan a disk and generate thumbnail images, store location, and can directly reopen the original image file. Many users may find this preferable. There are several useful image databases, including Multi-Ad Search 3.1 (Multi-Ad Services, Peoria, IL), Cumulus 2.5, Kudo and Kodak Shoebox (Adobe Fetch is no longer available). Of the various databases that I have tested, Multi-Ad Search 3.1 and Cumulus 2.5 are the most useful. Cumulus is particularly advantageous as it supports Apple ScriptMaker, and can automatically transfer thumbnail images to a FileMaker Pro database. A new low cost single-user version of Cumulus (3.0) is scheduled for release in the Fall of 1996.
7. Cataloging Software: Where Are The Files When You Need Them?
By this time, you will be overloaded with files, different disks, floppies, Zip, Jaz , magneto-opticals, Syquests and everything but paper tape. You will have various versions of the same file in many different locations. How will you ever find a file that you need for a publication?
Iomega Corp. (Roy, UT) provides a utility called "FindIt" with their Jaz drives that makes a compiled directory of the contents of various hard disks and removables. It works very well with magneto-opticals disks, Zip and Jaz drives, and floppies. However, it will only work with drives that have Macintosh formatting. It does not even recognize the presence of a PC-formatted Zip or PC-formatted magneto-optical disk (see below).
II. EVALUATING FLUORESCENT IMAGES WITH NIH IMAGE PRIOR TO CONFOCAL IMAGING
1. Evaluation Of Fluorescent Images Prior To Confocal Microscopy
The best way to obtain excellent final confocal images is to start with a good specimen. All the digital magic in the world won't make a good picture if you start with a bad specimen.
Careful evaluation of your fluorescent labeled specimen on a high quality fluorescent microscope will save you a lot of time and effort on the confocal microscope. Your best confocal images will be obtained from sections of highest quality. Such sections are easily identified on a standard fluorescent microscope. The qualities that should be evaluated include:
A) General tissue quality, including fixation, lack of tears or folding in tissue
B) High signal to noise ratio. The labeled processes should be bright and readily distinguished from the background. High background fluorescence poses a difficult problem that cannot be easily overcome by even the best confocal microscope. Poor signal to noise ratio will prompt you to modify your final image by increasing the contrast to excess.
C) Good separation of fluorophores. Very bright intensity of fluorescence of tetramethyl rhodamine isothiocyanate (TRITC) or indocarbocyanine (Cy3) will produce substantial breakthrough of the image into the range of the fluorescein isothiocyanate (FITC) image. While your naked eye will note this to be somewhat red in color, the PMT in the CLSM is colorblind and will detect this and not distinguish it from the FITC component of the image. While additional filters may help, there are several strategies that will help you deal with this problem.
- Do not use TRITC with FITC. Try rhodamine sulfonyl chloride (LRSC) or indodicarbocyanine (Cy5) in conjunction with an Argon-Krypton laser.
2. Fluorescent Excitation/Emission: A Moving Target
Fluorescent excitation/emission is a degradative process. The brighter the excitation (within limits), the brighter the resulting emitted image. In order to examine the image for adequate evaluation of its quality and content, the user will expose the tissue for increasing lengths of time. Both excitation brightness and duration of exposure, while required for evaluation of the tissue, also degrade the quality of the fluorescence. At saturating levels, the half life of FITC and TRITC is only about 1-2 seconds. When dealing with in vivo or in vitro specimens, the problem is even more notable, as the prolonged exposure to intense excitation light degrades the cells, and hastens cell death. (This has been used to good benefit in the selective killing of identified cells in nervous tissue). Achieving good dark adaptation by the observer, use of efficient microscopes, supercooling the specimen, pulsing the light source and various reagents (often toxic to living cells) have all been used to diminish the fading, but the inevitable lose of fluorescence cannot be fully avoided.
As a result of these limitations, you may find that you are reluctant to examine the tissue at length before going to the confocal microscope to capture your "perfect" confocal images. This results in many bad confocal images, often consequent to the fact that the original tissue was not properly evaluated before using the CLSM. All the tricks in the world of post-acquisition processing of confocal images won't improve a lousy specimen. An expensive CLSM won't take a good picture of a bad specimen.
Far too often, investigators would be better served learning how to optimize their use of a fluorescent microscope more thoroughly before spending time hacking images from a confocal microscope.
3. Selection Of Fluorophores
This is a large and complex topic. It is dealt with extensively in the excellent chapter by Brelje, Wessendorf and Sorenson (1993) in the monograph Methods in Cell Biology: Cell Biological Applications of Confocal Microscopy , edited by Brian Matsumoto.
If working with a single fluorophore, most users seem to prefer either FITC or Cy3. FITC has long been used, and the emission wavelength corresponds to the range of peak sensitivity of the human eye. Cy3, with an emission wavelength in the short red range, has become increasingly popular. Cy3 is intensely fluorescent and bleeds both high and low--use it only in single fluorophore configuration, unless you have previously determined that the two compounds of interest definitely do not co-localize in the same structure. The great advantage of so bright a fluorophore is that you can use a smaller aperture, with a favorable signal to noise ratio, lower laser power and shorter exposure times.
B) Double label fluorophores: FITC and LRSC or FITC and Cy5
C) Triple label fluorophores: FITC, LRSC and Cy5
D) Quadruple label: aminomethylcoumarin acetate (AMCA), FITC, LRSC and Cy5 (AMCA requires a laser that excites in the range of UV to short blue. While expensive, microscopes with lasers in this range are increasingly available.)
All these fluorophores can be obtained from Jackson Immunoresearch Laboratories, Incorporated (West Grove, PA).
4. Using NIH Image To Visualize Cy5
Many people have difficulty with Cy5 for the very reason that makes it so useful--it's emission is widely separated from that of FITC, and thus is barely visible to the naked eye. It is only visible if you are completely dark adapted, have a very efficient microscope and the intensity of fluorescence is robust. Even then, you will not be able to see much of the fine detail.
However, as users of NIH Image , you have one of the best tools available for imaging Cy5, if you have a video camera that supports on-chip integration (see following section) and a Scion LG-3 frame grabber. CCD video cameras are quite sensitive in the red to infra-red range. Wayne Rasband has provided a useful macro that allows you to vary the on-chip integration time and adjust the duration of integration while viewing the image on your computer monitor. For even greater control of signal fading, you should also use a Uniblitz shutter between the fluorescent light source and the microscope (see "Macros For Shutter Control" below).
5. On-Chip Integration
The intensity of fluorescent images is relatively low compared to the sensitivity of most CCD cameras. Video cameras sample available light, then send a complete image to the monitor every 1/30 of a second (33.3 msec or 30 frames per second). At that moment, the previous image is erased and the camera again starts to accumulate a charge proportional to the intensity of the incident light.
As the amount of light declines, the camera produces a more grainy, noisy image. With progressively lower light levels, the camera fails to detect a sufficient number of incident photons to provide a useful image. Averaging frames will not help, nor will software integration of multiple weak images.
In photography with film, the user has a number of options:
A) Increase the speed of the film
B) Increase the aperture or light collecting ability of the optics
C) Increase the exposure time--i.e., decrease the shutter speed
The analogous situation pertains to improving image capture with video cameras.
A) Increase the speed of the film
If you need to observe rapidly changing events, the only strategy you have available is to increase the sensitivity of your detector by using an intensified video camera. This is quite expensive, and the quality of the image is often degraded by the intensifier.
B) Increase the aperture or light collecting ability of the optics
There have been great improvements in the quality of fluorophores, light sources, lenses and filters. However, many images are still too faint to be adequately captured with a standard video camera.
C) Increase the exposure time--i.e., decrease the shutter speed
If your specimen is stable, then you can opt for longer exposure times.
On-chip integration acts by increasing the exposure time, thus increasing the number of photons captured. Rather than sending the image to the monitor every 33 msec, photons are allowed to accumulate on the sensor, and the image sent to the monitor when it is deemed adequately saturated. This technology has been available for many decades, but the cost often seemed beyond the budget of most labs.
NIH Image , in conjunction with some inexpensive hardware, now permits this technology to be widely used.
In order for this to work properly, you need four components:
A) A video camera capable of on-chip integration.
B) A source for a properly timed TTL pulse.
C) A frame grabber (video digitizer) that grabs the image at exactly the correct moment that the camera is sending the integrated image.
D) A computer and software that can control this process.
6. Practical Guidelines For Implementing On-Chip Integration
A) Choice of Camera: Several video cameras now provide built-in circuitry for on-chip integration at no additional cost. These include the two most widely used video cameras in labs, the Cohu 4915 (Cohu Inc., San Diego, CA) and the Dage-MTI 72 (Dage-MTI, Inc, Michigan City, IN). Images are allowed to accumulate on the camera sensor until a suitable level of exposure is achieved. A TTL pulse is then sent to the camera indicating that the computer now expects to receive the accumulated (integrated) image.
B) and C) TTL Pulse Source and Synchronized Frame Grabber: The Scion LG-3 card has made this technology available at low cost. Under normal operating conditions, the LG-3 continually samples the input from the video camera and displays the results to the monitor. However, under appropriate software control, the LG-3 card provides an output of a TTL pulse that is synchronized to the video digitizer, and captures the next image and holds it in the display buffer.
D) The software for controlling this whole process is provided in NIH Image . Wayne Rasband has provided a macro to control the duration of the exposure time for on-chip integration.
7. Macros For Shutter Control
I strongly recommend the use of Uniblitz shutter between the excitation light source and the specimen. The Uniblitz shutter is manufactured by Vincent Associates in Rochester, N.Y. Most microscope manufacturers can provide an adapter to mount the shutter between the excitation source and the microscope. The shutter is controlled by a Uniblitz controller box connected to the computer.
The advantage of using the shutter control is that it will limit the exposure of your tissue to the fading effects of the excitation by restricting exposure times to only that period required for actually collecting the images. Without this, I often forget to manually close the shutter, only to discover that I have burned out my best specimens.
Chi-Bin Chien, formerly of UCSD, has written a series of macros to control the shutter. I have combined the shutter control macros with Wayne's macro for on-chip integration. These macros are available as part of the collection of "Confocal Macros" provided at the NIH Image FTP site.
The following description pertains to the use of a fluorescent specimen. I suggest that you practice the use of on-chip integration with a transmitted light specimen, with the illumination set to a very low level. Once you have mastered the procedure for on-chip integration, you can then experiment with a fluorescent specimen.
A) A camera window should be selected. The shutter can be closed. Turn off the AGC (automatic gain control) on the CCU (camera control unit ). Set the manual gain and manual black level fully counterclockwise.
B) Select the macro for on-chip integration. If you do not have a Uniblitz shutter, you should open the manual shutter to allow excitation of your specimen.
C) If the image is too dim, move the mouse towards the top of the camera window.
D) If the image is too bright, move the mouse towards the bottom of the camera window.
E) The information window in the lower left corner of the screen will report on the number of frames that were integrated for the latest image. You can modify the macro to provide a direct readout of the duration of integration in seconds.
F) Observe the histogram for best spread of brightness values.
G) You may find it helpful to make minor adjustments in the quality of the image by changing the gain and black level settings on the CCU.
H) When you are satisfied with the quality of the image, terminate data collection by moving the mouse off the left side of the camera window and hold down the mouse button. The last image will continue to be displayed on the monitor. The shutter will then close, protecting the specimen from further fluorescent excitation.
This procedure requires practice to obtain images with a broad range of values.
As on-chip integration time increases, you will note a marked delay between the time you press down the mouse button and the appearance of a change in the image.
8. On-Chip Integration, Multiple Labeled Sections, And RGB Images
NIH Image permits you to evaluate the quality of multiply stained sections prior to using the confocal.
If you first obtain optimal quality images on the standard fluorescent scope using NIH Image , you can then combine the images to produce a useful RGB Stack. This can be tentatively evaluated using the NIH Image function RGB to Indexed Color . The Stack can also be saved and produces a high quality 24-bit RGB image that can be further modified in Adobe Photoshop.
A) Collect the image that you wish to be in the red slice of the Stack, using on-chip integration. Adjust the gain and black level to obtain the widest range of values, as indicated in a histogram of the image.
B) Run the macro provided "Make Stack from Current Image". This will take the current selected image, make a new Stack with three slices, for the R, G and B planes, and paste the selected image into the "red" plane.
C) Now collect a second image using the on-chip integration. Copy it to the buffer and paste it into the second slice ("green" plane).
D) If you have a triple labeled section, collect this with the on-chip integration routine, place it in the third slice ("blue" plane).
E) You can now use the RGB to Indexed Color operation under the Stacks menu to generate an indexed color image showing the double or triple labeled result.
F) Save the file as an RGB TIFF file. It can now be opened with Adobe Photoshop for better color images.
The steps in the above procedure can be further automated with a macro.
III. OPENING CONFOCAL IMAGES IN NIH IMAGE
The most commonly used CLSMs are made by BioRAD, Zeiss, and Leica instruments. The software/hardware used to generate images in these instruments stores the files in a DOS based file format. Thus the user is confronted with two initial tasks: transferring the file from a DOS based computer to a Macintosh and converting the file from its original file structure to an NIH Image file.
1. Transferring Files From A DOS, DOS/Windows Or OS/2 Based Computer To A Macintosh
I suggest that you initially collect your images to the hard disk connected to your system. This will speed up data collection of extended Z-series. At the end of each work-session you can transfer all the files to your Macintosh using one of three methods: A) Sneaker net, B) Local area network, or C) FTP via Internet.
A) Sneaker Net
Transfer the files from the storage location on the original disk to a Zip , Jaz , Syquest, magneto-optical or similar high capacity medium. These disks can be formatted as a DOS disk, and data directly transferred from BioRAD, Zeiss and Leica CLSM computers to these media. The Macintosh PC Exchange provided by Macintosh in System 7.1.2 and higher allows you to directly read PC-formatted floppies, Zip disks, and others. I don't recommend using Zip drives for original data collection as they are relatively slow, with access times of ca. 30 msec. The Jaz and Syquest drives have access/write times approximately equal to that of hard disks. For further comments and cautions, see below.
These disks can be manually carried to your Macintosh, which is presumably equipped with a similar drive. This is commonly referred to as a "Sneaker net". The major limitation of this method is that it demands that your Macintosh be able to read the various formats of different media. This is not always a valid assumption and relies upon your ability to find the correct drivers for different operating systems for the different types of media.
My own preference is for a disk of reasonable cost that can hold a typical workday's worth of data. The Zip drive, with storage capacity of ca. 100 MB and at a unit price of about $12-15, though relatively slow, meets both these criteria.
I don't recommend the use of floppy disks. They are very slow and cumbersome if you have many files, and requires that you have many pre-formatted disks. If you have an extended Z-series on a BioRAD CLSM, the file may be 7-25 MB, and is not easily transported via floppy disks.
Magneto-optical disks have a large capacity, and are long-lived. However, each drive manufacturer seems to have a different formatting scheme. I recommend them for archiving your files, but not for transferring them from the CLSM to the Macintosh. We have had endless aggravation with various incompatibilities.
B) Local Area Network (LAN)
An efficient method of transferring files is to have both the DOS and Macintosh based machines on a common Ethernet network, with a common local server. If you have an Office of Computer Services (or something of similar ilk), they can provide various options for LAN between Macintoshes and PCs. An alternate means is to use MacLAN 5.5 for your PC MacLAN (Miramar Systems, Inc., Santa Barbara, CA) operates within Windows 3.1 and Windows 95. MacLAN allows your Macintosh "Chooser" to see the PC as another client on an AppleTalk zone, and your PC to act as if it were another Macintosh on your AppleTalk network. Assuming that you are familiar with using a Macintosh on a Ethernet network, this allows simple file transfers at high speed. We have found MacLAN to be a good idea, but seems to be plagued with bugs and problems in reliability and speed of transfer of files.
C) FTP Via Internet
Transfer between computers that do not share a common server or network can be accomplished using Internet protocols such as FTP (file transfer protocol). In our experience, the most efficient program for doing this on the Macintosh is Fetch 2.1.2 (or higher), a public domain freeware program available on the zippy.nimh.nih.gov server. Fetch 2.1.2 will facilitate all aspects of the transfer from a PC/DOS machine. If you do not know how to set up your PC as an FTP site, contact your local computer center for assistance. The advantage of FTP using Fetch 2.1.1 on the Macintosh is that it doesn't seem to care what kind of operating system is working on the remote host. Fetch may be one of the most bomb-proof programs I have worked with--and it's free!
When performing an FTP, make sure that you perform the transfer in binary format, or the files will be unusable.
Do not erase the original files until you are certain that you have obtained a successful transfer and conversion. We recommend that you always save copies of the original files in their original native (DOS or OS/2) format.
A potential drawback to both LAN and FTP transfers is that the host computer containing the original files must be continuously available for the transfer. If someone decides to turn off the computer, or wants to use it for further data collection before you get back to your computer to effect the transfer, you may be cut off. In addition, if you are mainly a Macintosh user, you may find that setting up LANs and FTP links from the PC side is not as simple as on the Macintosh.
D) Zip Drives
The new Zip drives from Iomega have inexpensive removable media that hold 100 MB/disk. This is usually sufficient for a single day's CLSM collection. A major advantage of the Zip drive is that it is also compatible with the IBM OS/2. I have used this without difficulty with the Leica/Windows 3.1 version, BioRAD Comos for DOS, and LaserSharp 1024 running under IBM's OS/2. The advantage of the Iomega Zip drive is that the drive is small, relatively inexpensive, and is supplied in two different hardware versions that can be connected to either the standard SCSI port on the Macintosh and many PCs, or to the parallel port on PCs. Iomega provides drivers for use with MacOS, Windows 3.1, Windows 95, Windows NT and IBM's OS/2. The disks, if formatted for the PC, are also suitable for OS/2, and can be read on a Macintosh using the standard PC Exchange provided in the Macintosh operating system.
- Few Words Of Caution About Using Zip Disks
If you open a file on your a PC-formatted Zip disk on the Macintosh, and then Save it, you will have difficulty using the disk again on a PC. Iomega has not been helpful about the issue of cross compatibility of their drivers between PC and MacOS.
In view of the problems I have encountered with the Zip drives mentioned above, my current practice is to transfer the contents of the PC-formatted Zip drive to a magneto-optical with Macintosh formatting. If I want to use the files on a Macintosh (e.g., at home), I transfer the files to a Zip disk that has been formatted for the Macintosh.
The archival stability of Zip drives is still unknown. I strongly recommend that you store the original data files on archival media.
E) Magneto-Optical Disks For Archival Storage
The simplest means of storing large quantities of data is to store the file on a removable disk medium of large capacity that can be read by DOS, Macintosh and OS/2 systems. Since confocal images generate large data files, most systems have magneto-optical (MO) disks of 500 MB to 1.3 GB. A popular medium for this is the 1.2 GB MO disk, sold by Sony, Verbatim and others. These have the advantage of large capacity and have an archival life of at least thirty years. The most common drives are the Tahiti Max Optix 3, Pinnacle Sierra, various Sony, Ricoh, HP and NEC units. The disks are frequently (though not always) interchangeable. Many BioRAD confocal scopes were supplied with Panasonic MO drives. These use a proprietary disk that cannot be read by the drives of the previously mentioned manufacturers. When confronted with a Panasonic drive, use a LAN or FTP to transfer your files.
Until recently, these were the most common means of primary data storage for confocal microscopy. The price of the disks have dropped recently, and are now available for about $50 for a 1.3 GB disk. This is about half the price of a Jaz drive (also 1 GB), and though slower than the Jaz drives, is less prone to data loss.
The popularity of magneto-optical disks has declined recently due to the widespread availability and low price of Zip drives. Many people have had problems when attempting to read standard 1.2 GB disks that were originally formatted on a DOS based Pinnacle or Tahiti drive. This is particularly severe if you are using OS/2 with LaserSharp 1024. A significant problem with magneto-optical disks is the lack of universal driver standards. Use "Multi-Driver" on your Macintosh to facilitate reading DOS-formatted optical disks. "Multi-Driver" is a Macintosh Control Panel included in a software package sold by PC Access.
The BioRAD OS/2 optical driver cannot read Macintosh-formatted magneto-optical disks at this time. I no longer recommend using magneto-optical disks to move files from a PC platform (Windows or OS/2) to a Macintosh. Go with a Sneaker net using Zip drives, LAN or FTP. Magneto-optical disks, however, remain an ideal medium for archival storage of large files.
2. Opening BioRAD Files in NIH Image
BioRAD files consist of a 76-byte header that defines the size of the image in width and height, states if it is a Z-series, and, following the image data, provides information on parameters during collection of data, magnification scale and notes. The file structure of the new software, LaserSharp 1024, is similar to that of the earlier versions of Comos, with only minor exceptions pertaining to the footer at the end of the file that contains information about the parameters used during data collection.
Individual BioRAD files can be opened using the Import function of NIH Image . However, this requires that the user know the width and height of the individual file (Most frequently 768x512 in BioRAD files from an MRC600 512x512 or 1024x1024 in LaserSharp 1024 for an MRC1000 512x512 in Leica files). Unfortunately, I cannot figure out how to transfer a merged 8-bit BioRAD image to a Macintosh. The associated LUT (look-up table) is apparently stored in a manner or location that eludes me. If someone has solved this problem, please let me know. The Import function does not permit importing a Z-series. The macro for importing BioRAD files will correctly import a Z-series.
The macro "Import BioRAD MRC Z Series ", contained in the associated file "Confocal Macros", enables opening of either single sections or a Z-series. This works equally well on files generated with Comos and those generated with the recently released software, LaserSharp 1024. LaserSharp 1024 operates under OS/2. These files can be opened using the same macro employed to open the DOS-formatted files. LaserSharp 1024 stores each separate color plane of an RGB Z-series as a separate file. All three image planes of a single RGB file can be optionally stored as a single file.
After loading the BioRAD macro, you can "run" it by selecting it from the Special menu. The dialog will ask you for the "Starting Slice" of the Z-series. The default is 1.00. Accept that value for the moment. A BioRAD file containing only a single image will appear on the screen with the title of the original file (e.g., Axons03.PIC). I suggest that you immediately Save the file, prior to making any modifications, (e.g., "Axons03.PIC.Img"). This macro will load as many sections as possible, dependent upon memory.
If the number of Z-sections exceeds the memory allocated to NIH Image , the macro will stop running. You can Save the first group of sections in a separate file. Close the file, and again run the macro "Import BioRAD MRC Z Series", but when prompted for the "Starting Slice", enter the number of the last Z-section displayed. If the original Z-series is very large, you may have to do this several times. You can solve this problem either by buying lots of RAM, or store Z-series in smaller increments. This problem is particularly notable if you have collected images at 1024x1024 as well as collecting an extended Z-series. (e.g., 70 sections at 1024x1024 results in a file of ca. 74 MB.)
The associated calibration and notes of a single image file will be placed in a separate NIH Image text window. If you wish to include this information on the image that you saved as an NIH Image file, you must Paste it from the text window to the image window. If your notes contain information about pixel size and dimension of the image, use that to calibrate the scale of the image. (See "Set Scale" in NIH Image Manual ). Once you have calibrated the image, Save the file once again. The current version of the BioRAD macro does not properly read the calibration or notes file attached to a Z-series file.
BioRAD Z-series can be stored as a single large file, or as a series of individual sections. The former configuration has the advantage that all related files are stored in a single locus. However, it also means that such files may be huge. Using the above macro, a BioRAD Z-series will be opened into a new Stack. Save this in a similar manner suggested above for a single image file.
If this is a Z-series, use Stacks menu, and select Options to enter the step size of the motorized focus used when originally recording the Z-series. The magnification and slice spacing are stored with the original BioRAD file.
You may find it useful to Add a Slice to the beginning of a Stack to record specific comments about the series, what the data represents, and an image of Z-projection to demonstrate the contents of the file.
Save the original BioRAD *.PIC files, just in case. You can move all the original files into a separate sub directory to reduce clutter.
A) BioRAD Split Screen Images
Simultaneous collection of double labeled sections on the BioRAD can be displayed and saved on a "split screen" image, with the two images from PMT 1 and 2 displayed "side by side." This has two advantages:
- You can directly compare the location of different antigens simultaneously
The macro described in the previous section will open and display the split screen image. If you want to separate the two halves of the screen into separate images, use the macro "Merge BioRAD Split". This will convert the original split image into a three slice Stack (left image, right image and one blank black slice), then produce a merged color image. If you want to interchange the red and green planes, Select the window containing the Stack of three images, and run the macro "Swap Red_Green".
The Stack of three slices (left, right and blank), can then be saved as a 24-bit image in Adobe Photoshop 3.0 format, in the following manner:
- If you have run the preceding macro, and have generated an RGB image in NIH Image , NIH Image will save the file as an RGB-TIFF file. This format is accepted as an Adobe Photoshop 3.0 RGB image.
B) Merging Split Screen Z-Series
The macro that operates on a Z-series of a split screen multi-labeled section will:
- Place the left image in one Stack, and the right image in a second Stack.
Leica file format is a TIFF format. The Leica operating system is a peculiar hybrid using a VME bus with a 68040 CPU operating under OS/9, but uses a Windows front-end that generates DOS types of files. Leica has announced the release of a new software/operating system for their confocal microscope, based on Windows NT. The format of the images, however, will reportedly be unchanged. The original Leica files (and the new Windows NT files) can be directly read by a Macintosh. They will show up on your Macintosh desktop as PC files, with various optional icons, depending on how you set your system parameters in the Control Panel, PC Exchange . Each separate image file is accompanied by an "info.dat" file. Z-series are stored as single files with sequential numbers but with only a single info.dat file for the whole series. In order to reduce confusion in handling these Z-series with large numbers of files, move each Z-series set into a separate folder on your Macintosh.
If you are running NIH Image , you can Open these files directly, without having to Import them. However, you will not be able to double-click on the files to open them in NIH Image . In order to do that, pursue the following procedure:
A) Obtain a copy of "CTC 1.4" (or later) to change file type and creator. CTC 1.4 is available from the NIH Image server at zippy.nimh.nih.gov.
B) Select all the Leica graphics files in each folder (not the information.dat files) and drag them on top of the icon for CTC.
C) Using the resulting dialog box, change the "New Creator" to Imag , and the "New Type" to TIFF (creator and type are case sensitive, so enter exactly as spelled).
All the files will now have an NIH Image icon, and will be treated as NIH Image files by the program.
The info.dat files should be changed to "New Creator" Imag and "New Type" to TEXT . This file contains important information including scaling factors, size of image, size of pixel, and spacing in a Z-series.
To open a Leica Z-series Stack:
A) Close all other NIH Image windows at this time. Make sure that you have placed all the related Leica image files into a separate folder.
B) Select Open from the File menu. In the resulting dialog box, go to the desired folder and click "Open All", then Open . This will open all the files in rapid sequence.
C) Select Windows to Stack from the Stacks menu. Make sure that you set the magnification/calibration scales and slice spacing.
D) Save the resulting Stack with a new name.
4. Zeiss Files
Zeiss files saved using the newer version of software with the LSM 310 and 410 are "standard" *.TIF files. Use the same procedure outlined above for the Leica files--i.e., use CTC to change "New Creator" to Imag and "New Type" to TIFF . The files can then be directly opened by NIH Image . Make sure the LUT is correctly set. I have often found the Zeiss LUT map to be inverted, with a negative slope, and an inverted image. Click on the icon in the lower left corner of the mapping window to obtain the correct black/white relationship.
5. Molecular Dynamics/Sarastro
Jay Hirsh kindly provided information on the format of Molecular Dynamics files. The data files are simple files without a header or footer. Each image of a Z-series is stored as a separate file. Information about a single image, or about a Z-series is stored in a separate text file. The text file can be directly opened by NIH Image .
A) Place all associated images of a single Z-series in a single folder. Make sure that they are correctly numbered to reflect their order of collection (e.g., 008.ext, 009.ext, 010.ext, not 8, 9, 10).
B) Select Import from Stacks menu. The user must know the image format (512x512 or 1024x1024) in order to use this correctly. The offset value is 0.
C) Check boxes "Open All" and "Invert".
D) Click Open . This will open all the files to the screen.
E) Select Windows to Stack from the Stack menu.
F) Save the Stack with a name of your choosing.
6. Noran Confocal Files
There is a version of NIH Image that is used for direct data collection on the Noran. The resultant files are obviously compatible with NIH Image . The Noran is also sold with a version of Image-1, a DOS based program. A version of this program is now also available for DOS/Windows. I have no information about the file format used by Image-1.
IV. BASIC IMAGE MANIPULATIONS
NIH Image provides the user with an extensive range of image processing tools. You should become fully conversant with all these tools, including how to obtain a histogram, how to interpret the histogram, manipulating the look-up table (LUT), substituting a pseudocolor LUT in place of the standard gray scale LUT and inverting the LUT.
Keep in mind that most changes will only be made to the "display" buffer not to the "file" buffer. Thus, if you examine a histogram, then modify the brightness and/or contrast, the histogram will not be changed. If you now Apply LUT to the file buffer, the new histogram will reflect those changes. The original file, stored on your disk, will not be altered by this operation unless you now Save this modified file. The reader should use one of their own sample CLSM files to familiarize themselves with operations of NIH Image . All the operations in this section are fully dealt with in the NIH Image manual, About NIH Image .
You now can open one of the files that you imported to the Macintosh. Until you are experienced with the program, make a backup copy of the file, and only work on the copy, not the original file. Using the Save As. function of NIH Image , change the name of the file so that you don't mistakenly modify your original data file.
1. Evaluating The Quality Of Your Original CLSM Image
As stated previously, the quality of the image that you obtain from the following manipulations will be directly dependent upon the quality of the image that you start with. Thus, if you start with a lousy image, you may be able to make it look presentable, but it is still going to be a lousy image.
A) Making sure that your original CLSM image uses the full range of 8-bit values (0-255)
The most common flaw with confocal images, as with video images in general, is that the user has not utilized the full 8-bit range of gray values. Many of the manipulations that you perform on the original CLSM may make your image appear to be satisfactory. However, many data files do not contain a full dynamic range of values (0-255), and the image was massaged by artificially spreading a narrow range of values (e.g., 50-120) by modifying the LUT.
In order to develop an appreciation for the gray scale content of your images in NIH Image , examine a histogram of the image.
- Make sure that the image window is active by clicking anywhere within the window.
Learn to use the adjustments to black level and gain on the CLSM to optimize the spread of gray values. If you have to trade off between using the "+1 to +3 LUT" position on the BioRAD versus a brighter setting of the laser, choose the brighter laser setting. It will burn out your specimen faster, but give you a better signal to noise ratio and a wider dynamic range in gray scale value. The "Photon Counting" mode, or the "Accumulate" mode on the BioRAD will often provide an image with the best dynamic range. On the BioRAD, I prefer to use the "Accumulate" mode with "Slow Scan" for best results. It is not the purpose of this manual to teach the use of CLSMs, but I only wish to emphasize the importance of the quality of the original image.
B) Avoiding High Contrast Images
If you get into the habit of checking the histogram of your images as you collect them on the CLSM, you will improve the quality of the original images, and find less need to twiddle with the LUT values.
C) Avoiding Noisy Images
If possible, collect images using the "F1" (Slow) setting (on the BioRAD) with a Kalman setting of at least 3, or "Accumulate" mode. When using "Accumulate mode", you can use a less intense laser source, and manually accumulate until the image appears satisfactory.
2. Editing Image
A) Cropping Images, Erasing, Superimposing Text, Scale Bars, Rotating And Scaling Images
NIH Image provides a wide range of tools for editing your image. These are described in NIH Image manual About NIH Image .
- Notes On Scaling
- Caution On Scale And Rotation Of Images
3. Using LUTs
The following operations are commonly used to enhance the image. These are standard operations on all confocal scopes, and are based on methods that are widely used for manipulating digital images. See appropriate section of the NIH Image manual.
A) Modifying Brightness And Contrast
In the simplest operation involving look-up tables, you may choose to emphasize a selected range of index values (gray scale values), and minimize other values. The choice may be based on your interpretation of the information content of the image. Learning how to interpret the histogram, and modifying brightness and contrast are essential skills in image processing.
B) Linear And Non-Linear LUTs, Including Custom LUTs
The standard LUT shown in the mapping window (lower left side of your screen) is a linear LUT. You can modify the brightness and contrast values by either dragging the Brightness and Contrast slider buttons, or by directly dragging the plotted line in the mapping window.
The "Confocal Macros" file included with this manual provides a number of alternate, non-linear LUTs based on sampling the values within your display buffer. These include Log, Parabolic, Square, Square Root and Gamma transforms. Play with each of these and observe the effects on your images. You will probably find the Gamma transforms most useful when used with a setting of 1.5 to 2.0. Many of the resultant changes produce results similar to those obtained on the BioRAD with +1, +2 and +3 LUT settings. When you open BioRAD files with NIH Image , the results may not match what you saw on the BioRAD. Most commonly, the image will appear much darker, and lacking the detail you so clearly remember seeing on the screen when collecting the original image. NIH Image has not corrupted your files. This is most frequently due to the fact that the image you viewed on the confocal microscope may have had a non-linear output LUT attached to it. The original data is imported intact, but the output LUT is not attached to the new NIH Image file, and a new linear LUT is appended to the file. See the NIH Image manual for instruction on how to modify the LUT.
C) Enhance Contrast Operator In NIH Image
The Enhance menu item in NIH Image has a specific function entitled Enhance Contrast. This produces a custom linear LUT effect that is as good as any result I can obtain in my attempts to manually modify the LUT curves. However, you may find that this operation produces an excessively contrasty image, with too steep a slope in the LUT mapping window. If so, then manually change the LUT in the mapping window to give a slope of the LUT about halfway between the original value and that produced by Enhance Contrast . If that is satisfactory, then Apply LUT (Command+L ). If you feel that you want still more contrast in the image, repeat the above sequence.
In order to appreciate the effects of this operation on the original image, examine a histogram of the image before and after this procedure.
As in all alterations to the LUT, the Enhance Contrast operation only modifies the display buffer, not the image in the file buffer or the original file on the disk. In order to alter the file buffer, you must perform Apply LUT . This will not alter your original disk file. If you wish to do so, Save the file. Once you have done that, you cannot go back to the previous image. You may, therefore, prefer to save the modified file under an alternate name.
Enhance Contrast is very different from the Equalize operation, which is likely to result in excessively splotchy images. Compare the effect of each of these operations on the appearance of the mapping, histogram windows and LUT.
D) Thresholding And Density Slicing
See NIH Image manual.
E) Pseudocoloring Images
Pseudocoloring confocal images assists the viewer in displaying double labeled sections, detecting major differences in concentration, or changing concentrations, as in calcium ratio imaging. See appropriate menu item, and NIH Image manual for use of this operation. All the color applications described in this manual, although they may resemble the original colored fluorescent image, are pseudocolor.
F) Exporting To Adobe Photoshop
Recent versions of NIH Image save single files and RGB files in a format that can be directly opened by Adobe Photoshop 3.0. This will not work with Z-series.
4. Enhancing/Filtering Image
NIH Image provides a wide range of tools for enhancing and filtering images, including filters to sharpen, smooth, shadow and detect edges. You can write your own kernels, run median filters, and Sobel operators. Some of these operations are multistage operations and alter both the display buffer and the file buffer (but not the disk file), and cannot be Undone . You will have to reopen the original file to restore the image.
For further details, see NIH Image manual.
5. Quantitative Measurements
NIH Image was originally developed for quantitative measurements. There are many useful functions described in the About NIH Image manual. Various measuring functions include: length, area, density, particle counting, and profile of a Line. About NIH Image will provide guidance in the use of these operations.
6. NIH Image Macro Language
One of the most powerful features of NIH Image is the macro language. This allows the user to write simple Pascal-like scripts. The distribution kit provided from the FTP site contains many sample macros that can be readily modified to your particular needs.
V. Advanced topics
1. Merging Pairs Of Double Labeled Sections
One of the most valuable and commonly used features of confocal microscopy is the ease of obtaining images of double and triple labeled histological sections. There are two major obstacles that may cause difficulties when attempting to merge two images:
A) The contrast and brightness values of one section may be markedly different from the other(s). This is best dealt with by careful evaluation of the images at the time of original data collection, and modifying your means of collection.
B) The two series of images of such a pair may not be in perfect register with each other. This may be due to various factors, including misalignment of the PMTs, the mirrors, and filter blocks. Most of the errors appear to occur in translation (X- and Y-axes) and not in rotation. A shift of a few pixels may not be noticed, but occasionally the error results in a marked shift from one color plane to the second. Simple translation errors can be corrected by shifting the images one or more pixels at a time. Rotational errors are more difficult to correct, take longer, and frequently result in image warping. If you find that you have marked rotational errors, the service personnel from the manufacturer of your confocal microscope should deal with this.
NIH Image provides an alignment operation, "Register". This can be used on sections in a Stack (see below).
A) NIH Image And Adobe Photoshop
Two or three gray scale images of different fluorophores can be combined into a single colored image, with each fluorophore represented by a different color. The color chosen to represent each fluorophore is arbitrary, and can differ from the original one. There are two different programs that can be used successfully for this purpose, NIH Image and Adobe Photoshop. There are benefits and disadvantages in the use of each program. Adobe Photoshop is relatively expensive, but a superb commercial program. It supports 24-bit images and allows almost instantaneous adjustment of the individual red, green and blue planes of a merged image. NIH Image produces an 8-bit custom palette of the merged image. It takes 5-10 seconds to produce this image on a Macintosh IIfx, and is correspondingly faster on a Quadra 950 or a PowerPC. Although this is an 8-bit color image, the result is often satisfactory, and occasionally even comparable to that obtainable with Adobe Photoshop. However, NIH Image may produce excessive dithering of the resultant image, and you cannot make small adjustments in brightness or contrast of the final color image obtained with NIH Image . Instead, you have to go back to the original gray scale images, modify them, and then once again Merge the images.
Since each "Indexed Color" image produced with NIH Image has its own unique LUT, you cannot directly do a side-by-side comparison of two different color images if you merged using "Custom Colors", as the color values shift markedly as you change windows. If you selected "System Colors", the quality of the color is more limited, but you will be able to compare results with other windows merged using the same system LUT. This is a major disadvantage in relying exclusively on NIH Image . However, for routine operations, NIH Image is satisfactory.
In comparison, Adobe Photoshop, using a full 24-bit window, allows you to compare multiple colored images simultaneously on a single screen. Adobe Photoshop also provides an excellent range of filters and convolutions. The more immediate advantages of NIH Image are manifest in measurement capability, generating Stacks, Z-series projections, and 3D-projections and rotations. Adobe Photoshop does not provide such facilities.
Until recently (prior to version 1.56), NIH Image could only run under an 8-bit monitor setting. If you wanted to shift back and forth from NIH Image to Adobe Photoshop, you had to reset the monitor to 24-bit. Beginning with version 1.56, NIH Image operates satisfactorily with the monitor set to 24-bits.
NIH Image version 1.59 now directly allows the user to save an RGB stack of three sections in a format that can directly be read by Adobe Photoshop 3.0. If the file is modified in Adobe Photoshop, then saved as an Adobe Photoshop TIFF file, it can be re-opened by NIH Image as a three slice stack. However, if you add additional Layers or Channels in Adobe Photoshop, it forces you to Save the file in Adobe Photoshop format. This cannot be read by NIH Image . An NIH Image Z-series containing Merged color slices cannot be read by Adobe Photoshop.
B) Double Labeled Sections: Building A Stack (Best Done Using A Macro)
- Color Merge In NIH Image
Two color plate: If you wish to combine only two plates, open the two files.
Use the macro "Color Merge Two Images" contained in the sample "Confocal Macros" that is provided with this manual. If you examine the sequence of commands in the macro, you will find the following operations:
A) Open a fresh stack
B) Paste the "red" image into the first slice
C) Add Slice to add an additional slice
D) Paste the "green" image into the second slice
E) Add Slice to add a black (empty) third slice
F) Merge from RGB to 8-bit color using a custom LUT
G) Open a new window containing the merged color image
Alternately, you can do this operation manually to familiarize yourself with the procedure. The first file opened will become the red plane, the second file the green plane. Under menu item Stack choose Windows to Stack . This will put the two plates into a stack labeled "RGB". Any further operations will not alter your original data files, so you can always go back and start again.
Save a copy of the NIH Image Stack. The following section describes how to use this Stack with Adobe Photoshop 3.0.
If you want to alter the contrast or brightness of either of the color planes, you have to modify either the original images or those in the RGB stack. I suggest that you limit your initial attempts to the slices in the RGB stack. When you change the Brightness or Contrast of any single image, you must then Apply LUT (Enhance menu) to that single image. Do not use the Apply LUT to "Stack" macro as this will modify all the images in the Stack.
From the Stacks menu select RGB to 8-bit Color. In the resulting dialog box, select "Custom Colors". This will generate an "Indexed Color" window from the Stack, but will not alter the stack itself.
Since each original file may have a different distribution of values (see histogram), the saturation of each color plane may differ markedly. Be creative, try different ways of doing it. (e.g., use Enhance Contrast operation on one slice at a time). Try non-linear LUTs, various filters, etc.
The resulting "Indexed Color" image can be saved with a unique name.
Once you have mastered this sequence, you will have a clearer understanding of the operation of the macro "Color Merge Two Images" contained in the sample "Confocal Macros" that is provided with this manual.
If you now Save the RGB stack (RGB TIFF) in NIH Image , the resulting file can be viewed with Adobe Photoshop as a 24-bit file.
- Merging Files With Adobe Photoshop
Version 1.58/1.59 of NIH Image now permits Stacks to be directly saved in Adobe Photoshop 3.0 TIFF format.
A) The stack must consist of 3 slices.
B) Before saving the stack, open Slice Info in the Stacks menu. Confirm that RGB is selected.
C) Save the file. It will write the file as an RGB TIFF file.
You can then directly open this NIH Image Stack in Adobe Photoshop 3.0.
An alternate means of converting the stack to an RGB TIFF format is to call the RGB to Color function. This will automatically change the "Slice Info" window to RGB. The resulting "Indexed Color" image will give an approximate (8-bit) preview of the results to be obtained with Adobe Photoshop (24-bit). Save this file.
You will now be able to use the various Adobe Photoshop tools to modify the resulting 24-bit image. You should explore the use of the Levels command (Command+L ), as well as the Brightness/Contrast command (Command+B ).
If you have a quadruple (or more) labeled section (e.g., 512x512, four fluorophores, or three fluorophores and a Nomarski/DIC image), and have stored them in a four slice stack in NIH Image , you can open them in Adobe Photoshop using the method described below.
A) From the Adobe Photoshop File menu, select Open.
B) Using the Adobe Photoshop Open dialog, go to the desired directory containing the NIH Image Stack file of interest.
C) Using the same dialog box, pull down list at the bottom of dialog box, choose Raw file format.
D) This will show all files in the chosen folder.
E) Select the NIH Image Stack containing the four slices. You have to know the dimensions of the slices.
F) The resulting dialog box requests that you fill in:
H) Save the new Photoshop image using the Save As. dialog box. Do not save as a Raw image. Select TIFF format, and choose a new name for the file, otherwise the simple Save command will overwrite the original NIH Image file and store the data in the Raw format.
You can modify each color channel separately using several different methods, as described in the Adobe Photoshop 3.0 manual.
The quality of the resulting color image will generally be much better than that provided by NIH Image , as it can utilize a full 24-bit look-up table, and does not require dithering of the image.
The Adobe Photoshop manual and tutorial will provide further guidance in modifying the images.
- Preferred Method
C) Compare Results Of NIH Image 8-Bit Merge With Photoshop 24-Bit Merge
The quality of the color image (assuming your monitor is set for 24-bit color) is generally much better in Adobe Photoshop than the optimized 8-bit image obtained in NIH Image .
D) Adjusting Color Contrast On Sections In A Stack (See Macros)
Do the Enhance Contrast operation separately on each section in the stack. Failure to do so will result in the merging of both saturated and unsaturated images.
If you are dealing with a single RGB section, you will find that Adobe Photoshop provides much better tools for this.
E) Merging A Double Labeled Pair Of Z-Series Using A Macro
- Open the two Stacks. The first stack will be assumed to be the "red" stack. The second stack will be the "green" stack.
The drawback of this procedure is that the resulting images are 8-bits, not 24-bits. I cannot find a way to do this procedure with Adobe Photoshop.
2. Projection Of A Z-Series And 3D Rotations
Many of the most valuable qualities of confocal Z-series are that the data can be manipulated to obtain "through views" of the stack, the stack can be resliced at various angles converting a transverse view into a sagittal or horizontal plane, it can be used to generate 3D images, stereo pairs, and other operations. Indeed, NIH Image is able to perform operations as well as much of the software provided by the manufacturers of confocal microscopes, and equivalent to many of the basic operations provided by very expensive image processing programs such as VoxelView, VolVis and VoxBlast running on a Silicon Graphics Workstation. For more elaborate operations using true Voxels, complex shading and ray-tracing in 24-bits, these latter programs are excellent choices. For the most common operations, however, NIH Image is quite adequate. Optimal performance on these tasks will be achieved using a PowerPC with sufficient memory to handle your typical data sets.
A) Stepping Through A Z-Series Using Stacks
- Open a Z-series in a Stack.
B) Animating A Z-Series
- From the Special menu, select Video for oscillating movies. This will produce smooth back-and-forth motion of the Z-series animation.
This is an extremely useful benefit of the confocal microscope. A stack of Z-series sections are all in optimal focus. If projected onto a single plane, all objects throughout the thickness of the imaged section will now be in sharp focus and spatial relationships may be more evident.
NIH Image version 1.58 and later provides macros to perform the Z-projection. A sample macro to accomplish this operation is included in the accompanying macro files.
- Selection Of Area For Z-Projection
A) Choose an extended Z-series (e.g., more than 10-20 images) with well defined profiles of objects.
B) Run the macro "Project Z Series".
C) Use Project command
D) Set :
Slice Spacing (Pixels): Set with your slice spacing
Initial Angle (0-359): 0
Total Rotation (0-360): 0
Rotation Angle Increment: 0
Lower Transparency Bound: 0
Upper Transparency Bound: 100
Surface Opacity (0-100): 0
Surface Depth-Cueing (0-100): 100
Interior Depth-Cueing (0-100): 0
E) Select "Minimize Window Size"
F) Select "X-Axis"
G) Select "Brightest Point"
After you develop a sense of familiarity with the program, play with different values. Start with the default values. Once you are more familiar with the Thresholding tool (Magic Wand), you will be able to use that to set the range of values desired for your Z-series.
This will generate a single Z-projection of all the plates in the Stack, viewed from directly in front of the stack.
Experiment with different view angles by using different values for the "Initial Angle", while leaving "Total Rotation" at 0.
If you want to make a quick and dirty stereo pair of images from a Z-series, then the values should be something such as:
Initial Angle (0-359): 356
Total Rotation (0-360): 8
Rotation Angle Increment: 8
Select the "Y-Axis" as your "Axis of Rotation". This will cause the resulting image to rotate left to right (i.e., around the Y-axis). Choose "Brightest Point" as the "Projection Method".
This will produce a new Stack containing two images at an 8 increment. This is a common angle for stereo pairs. In order to visualize this, you can do two different procedures.
- From the Stacks menu, Animate the stack, and the image will rock back and forth around the Y-axis.
A) Make sure that "Invert Pixel Values" are not selected in the Edit/Preferences menu item. If selected, de-select this item. Then Select File/Record Preferences. Quit NIH Image in order to implement the changes in preferences. If this item is "selected", you will be easily confused by the various values displayed in the info, mapping and histogram windows.
B) There are a number of parameters in the Project item under the Stacks menu. Understanding the proper use of this item is critical to obtaining pleasing results in Z-projections and rotations.
C) The first of several items include obvious settings regarding "Slice Spacing", "Initial Angle", "Total Rotation" and "Rotation Angle Increment". These values are all obvious and pertain to the sampling interval of the resulting projection.
The remaining items on the list, "Lower" and "Upper Transparency Bounds", "Surface Opacity", "Surface-Depth Cueing", and "Interior-Depth Cueing", however, are very confusing, and cannot be easily understood without some explanation.
One of the greatest difficulties in mastering these functions is because once you have selected specific settings, obtained an image, and then try other settings, your screen will be cluttered with images, and the particular parameters used to obtain them has been forgotten.
Start with the "Lower" and "Upper Transparency Bounds" in relationship to the stereo pairs you generated a few minutes ago.
When making a simple single view Z-projection, the slice spacing is irrelevant. However, depending upon the setting of the "Lower" and "Upper Transparency Bounds", and "Interior Depth-Cueing", or the resulting image is likely to appear somewhat lacking in brightness.
"Lower Transparency Bound" must be set to 0. (Note that this value is expressed from 0-254). This value determines the cut-off point of displayed pixels. All pixel values below the setting selected will be discarded and modified to a darker value. Remember the whitest/brightest pixel on the Macintosh has a value of 0, the black pixel a value of 255. "Lower Transparency Bound" greater than 0 (e.g., n ), will modify all pixels from zero to n in your image and replace them with darker pixels. This results in dark gray to black holes in the midst of a bright object. The setting of this value is quite simple. The most effective way to appreciate the consequences of this choice is to look at histograms of the images produced with "Lower Transparency Bound" set to zero, and compare that with a histogram generated with a "Lower Transparency Bound" of perhaps 50. Make sure that your starting image has prominent objects with histogram brightness index values of 1-10 (out of a range of 0-255).
For the "Upper Transparency Bound", I suggest that you start with a value of 254. After using this for a while, play with other values.
The three following functions in the Project dialog box are expressed as values of 0-100%.
Surface Opacity (0-100): 0
Surface Depth-Cueing (0-100): 100
Interior Depth-Cueing (0-100): 0
NIH Image 1.61 sets "Interior Depth-Cueing" to a default of 50. This will produce "muddy" images. You must set this to 0.
D) Reslicing The Z-Series Along Alternate Planes: (X-Z, Y-Z And Theta-Z)
One of the most useful features of NIH Image is the ability to rapidly reslice a Z-series data stack along alternate planes. In addition to simple orthogonal planes (i.e., X-Z and Y-Z), you can reslice at any arbitrary angle between X and Y to generate a single Z-plane cross section. This is very useful for evaluating the extent of penetration of antibodies into tissue, evaluating cell spacing, etc.
The closer the images in the original Z-series, and the greater their number, the more natural appearing the final results. In order to provide a seemingly continuous image in the resliced plane, the software interpolates gray values. For improved quality of resliced images, your original Z-series should be as close as possible (e.g., 0.25 m).
Have you set the calibration for magnification and slice spacing, as described above? (Also see NIH Image manual).
Your original data file may contain the needed information about slice spacing. In many of our files, the step size was 0.38 m (on a BioRAD), and the pixel size 0.105 m (9.5 pixels per m). However, this depends upon the Z-axis step size, the objective and the zoom factor.
A) From the Stacks menu, select Options . Enter the appropriate value in the slice spacing dialog.
B) Select an image from the series that best shows the features of interest. Using the Calibration/Measurement tool (dashed line in the right hand column of Tool Palette), draw a line across the section along the desired plane of resectioning. If you wish to constrain the plane to either the X-or Y-axis, press the Shift key and hold down as you draw the line.
C) From the Stacks menu, choose Reslice (Command / ). If circular profiles appear excessively flattened in either the X- or Y-axis, experiment with different values of "Slice Spacing" in the Options dialog.
E) Reslicing To Make A Z-Series In An Alternate Plane
The preceding instructions describe how to obtain a single reslice in the X-Z or Y-Z plane. If you would like to obtain a new stack of Z-series section, not just a single section, use the macro provided, "Reslice Horizontally" or "Reslice Vertically". The reslicing is limited to orthogonal planes of section. Thus, if you wish to reslice along an oblique angle, you will first have to rotate the stack. In order to do this, use the Angle Measurement tool from the Tool Palette. Measure the degrees of variance of the object in the image from the angle you finally desire.
A) Use the macro "Scale and Rotate Stack" , using the angle value established in the previous paragraph.
B) Now select a rectangular area that encloses the region you want to reslice in either horizontal or vertical plane.
C) Use the macro "Reslice Horizontally" or "Reslice Vertically".
1. Rapid Dynamic 3D Reslicing
Norbert Vischer of the Netherlands has recently contributed an extremely useful macro, "3D Slicer" for reslicing a stack of sections along the X-, Y- and Z-axes. This macro is provided in the program "Object-Image 1.59", available from the NIH Image FTP site. Object-Image 1.59 provides extremely rapid reslicing in real time. The individual resliced sections cannot be saved, for unlike the macros described above, they are not placed in a new stack. The reslicing of sections is accomplished by dragging the mouse along the X-, Y- or Z-axes of a master stack, and results in rapid generation of resliced images in the two other planes. Reslicing can only be done along orthogonal planes.
A) Open a Z-stack of sections, such as the sample MRI Images of a human skull and brain.
B) Enter the correct magnification scale and slice spacing (5.0 mm), as described above.
C) Under Stacks menu, select 3D Slicer. This will open a new window containing an angled perspective view of the original stack, with 3D-projection planes along the two alternate planes, as well. In the upper part of the window will be an image of the resliced section parallel to the X-axis. On the right side of the window will be the resliced section parallel to the Y-axis.
D) Place the mouse over the X, Y or Z margin. The cursor changes to a two headed arrow indicating the direction of movement. Drag the X-, Y- or Z- axis on the main central image, and observe the rapid resectioning in the alternate planes.
E) Place the mouse over the intersection of the X- and Y-axes and the cursor becomes a four-pointed arrow. You can now simultaneously reslice parallel to both the X- and Y-axes.
F) Double clicking the mouse on the X-or Y-axes will turn off the reslicing tool for that axis. Double clicking on the dashed lines on the X- or Y-axis will restore the reslicing planes.
F) Generating A 3D-Series From A Z-Series
This is a computationally intensive procedure. A fast PowerPC will prove very desirable when doing this operation. When you are still learning the basics of this procedure, use the Selection tool from the Tool Palette and outline a small portion of the image when generating a 3D-series. It will complete the task much more rapidly.
Use care in settings. Remember to correct for thickness and spacing of individual sections.
- Select Project from the Stacks menu.
Initial Angle (0-359): 0
Total Rotation (0-360): 15
Rotation Angle Increment: 180
Select the "Y-Axis" as your "Axis of Rotation". This will cause the resulting image to rotate left to right (i.e., around the Y-axis). Choose "Brightest Point" as the "Projection Method".
Now click OK and wait. The program will generate a new stack of images of the "rotating" objects in the original stack.
Once you have had further experience with this procedure, Project using a full 360 degree rotation at closer intervals, changing transparency values, axes of rotation and the various other options available.
A macro to facilitate this operation is provided.
G) Animating A 3D-Series (Producing "Apparent Rotation")
Under Special menu, select Video Options . Check "Oscillate Movies". Close the dialog box.
To generate the impression of rotation with your newly generated 3D stack, use the Stacks menu and select Animate (Command+= ). This is heavily memory dependent, so keep your initial efforts small.
You can control the speed of rotation with keys 1-9. You can step through single sections using the < and the > keys.
H) Make A Stereo Pair Or Series Of Stereos?
There are several methods of generating stereo images from a stack of sections collected in the Z plane. One simple method, the "pixel shift" method, is that used in the original BioRAD software.
The pixel shift method starts with a stack of a Z-series. You will generate two Z-projections from this original stack. One will be "pixel shifted" to the left and a second "pixel shifted" to the right. This is achieved by shifting each successive section in the stack by one additional pixel (or more, if desired) prior to performing the Z-projection. The two resultant images are then placed side by side. The pixel shift method is generally limited to stereo images centered around the original plane of image collection.
Projecting and ray-tracing is a second method that projects (ray-tracing) the stack onto an imaginary view plane from different angles of view and generating a 3D rotation series. This second method is computationally more complex, but provides the possibility of generating stereo views from any angle around a central point.
Start with the original Z-series Stack. Select the Project from the Stacks menu and set "Initial Angle" to 354, "Increment Angle" to 6 and "Total Rotation" to 12. This will produce a new Stack containing three slices (354 , 0 and 6 ).
1. Stereo Series In Black And White
Once you have generated a 3D-series, make a Montage series using the appropriate function from the Stacks menu. The Montage function is non-destructive, i.e., it does not erase the 3D stack, however, I suggest that you Save your work as you go along.
A) Stacks Menu: Montage
The resulting dialog box will show a series of values that are dependent upon the number of sections in the stack. If you made a simple stack with only 3 sections, then choose 1 row and 3 columns, and an increment of 1. If you generated a full 360 rotation series, you can select any range of slices (e.g., slices #4-8 out of 16 slices), or incremental slices (every 2nd or 3rd slice in the stack), and define the numbers of rows and columns you wish to see displayed.
This will generate a series of images in a new window on the screen. If the resulting images are too large to line up next to each other, use a scaling factor when generating the montage.
Start with two side-by-side images. To facilitate seeing the stereo effect without special viewers, choose a set of images with a prominent object to provide an alignment cue in the center of each image. Set the image size and separation with the alignment cues no more than 50 mm apart on the screen. Gradually work your way up to greater separations and then slightly larger images. Initially, you may find that you require a stereo viewer to visualize a stereo image from side-by-side images. With practice, you will not require a stereo viewer and should be able to scan across pairs of sections and jump from one stereo image to the next. (The typical interpupillary distance of most people is about 65 mm. You can also practice learning to fuse stereo images by using some of the recent popular books with "random dot stereograms", such as The Magic Eye .)
2. Stereo Pair Of Single Labeled Section In Color
An alternate manner of presenting stereo images is to merge a stereo pair into a single image plane, assigning one image to red and the other to green.
To generate a 3D rotation series around the Y-axes., limit the number of image to two at a separation of ca. 6-12 . For the first trials, set the "Initial Angle" to 354, the "Increment Angle" to 6 and the "Total Rotation" to 12. This will generate three plates in a new stack. Animate the new stack of images, and if the animation produces the desired effect, delete either the middle or one of the end plates. Add a black plate as slice 3/3. (See section above on Merging Slices to generate 8-bit color images). Using the Stacks menu item RGB to 8-bit Color , you will obtain a single merged image which can be viewed with a pair of Red/Green stereo glasses.
The greater the separation between plates (i.e., 12 rather than 6 ) the more exaggerated the stereo effect. Play with different values. Some people find that angles greater than 15-18 are excessive.
3. Color Stereo Images Of A Double Labeled Section
Although this is a more complex set of operations, it is a logical extension of the methods described above.
First generate a 3D rotation series of each Z-series of a double labeled section. Save the results. Close all windows except those of the two 3D rotation series. Now use the macro "Color Merge of Two Stacks". You will now have a rotation series (3D) of two simultaneous different fluorophores. Animate the series.
Now generate stereo pairs, using the montage method described above.
I) Exporting Stacks to QuickTime Movies and VCR Recordings of Stacks
A rotating Z-series, or any other Stack, can be saved in QuickTime format, allowing it to be viewed with a variety of other programs, such as a simple QuickTime Viewer, or with a program designed for preparing and transcribing clips on a VCR, such as Avid VideoShop 3.0. The latter program is presently distributed at no additional cost with many Apple Computers. Avid Videoshop can be used to prepare a VCR tape of your data, merging different data sets into a single sequence, etc.
- Saving Stacks In QuickTime Format
A) Save the Stack in standard NIH Image format.
B) Open the Stack, and select Save As.
C) There are a series of optional buttons at the bottom of the dialog box, including TIFF, PICT, PICS and others.
E) Rename file with new name (e.g., if file name was "NewCell", rename it "NewCell.PICS") to avoid overwriting your original data file.
F) The new file can still be opened as a Stack within NIH Image .
G) If you Save the file, it will revert to the original TIFF format, though with the new filename, and is likely to be confusing when you try to open it with a QuickTime player. Therefore, any additional Save operations should always be done using Save As. .
1. Video Printer
You can obtain an instant print of the NIH Image window using a video printer, such as the Mitsubishi 67U or Sony thermal video printer. This provides a gray scale print. The Mitsubishi produces a print of approximately 3x4 inches, and provides a running record of your results. Scion markets a NuBus board (TV-3) that will directly send the contents of a selected window to an NTSC device. The driver for the TV-3 board must be placed in the NIH Image Plug-Ins folder. The drawback of the TV-3 is that it can only send out an image of maximum dimension of 640x480 pixels. Since the standard BioRAD image is 768x512, the image must first be scaled to a 640x480 size, or select a portion of the image corresponding to this size. Images smaller than 640x480 will only fill a portion of the printed field. It would be useful if the driver provided an autoscaling function. A simple macro, however, can be written to accomplish a similar result. The new PowerPC 8500 provides direct NTSC and S-Video output. I have not had a chance to test them with the Mitsubishi printer, but based on published specifications, these should provide results similar to those obtained with the Scion TV-3 board.
2. Dye Sublimation Printer
Prints of much better quality, including black and white or color prints, can be obtained using the Kodak 8600 dye sublimation printer. The printer is provided with a Adobe Photoshop Plug-In driver. Place this Plug-In in the folder of the same name in the main NIH Image directory. When you want to print the contents of a window, select the desired image. Under the File menu, choose Export , and select the "Kodak 8600" printer driver. The typical image will print in approximately one minute. If you are in NIH Image , this works only with monochrome images printed on a black ribbon. Color prints are best produced using Adobe Photoshop. A major limitation in the use of the Kodak printer is the cost of each print ($2-$3), as well as the cost of the printer itself (ca. $8,000).
The wider availability of color laser printers may provide satisfactory color prints at lower cost than those obtained with the Kodak 8600 printer.
3. Slide Maker
There are a number of slide makers, including the LFR Lasergraphics, the GCC/Polaroid slidemaker, Agfa and others.
If you are using Microsoft PowerPoint to send images to a slidemaker, Scale the images to the full size of the screen (e.g., 768x512) prior to saving them. Use the "Bilinear Interpolation", rather than "Nearest Neighbor", for best results. You can also use PowerPoint to combine different images on a single slide, add text, or other graphics elements. Paste the NIH Images onto a black background to avoid having a white border around the image.
Videotape players are now commonly available at all universities and scientific meetings. You may find it far more effective to display rotating projections as well as sequences of sections on videotape, rather than with 2x2 slides or overheads. The technology for transferring such Stacks is widely available, and a brief description of its application to NIH Image and confocal Stacks is described in the preceding section.
VII. USE OF NIH IMAGE FOR IMAGE COLLECTION AND INSTRUMENT CONTROL
The majority of CLSMs have their own dedicated software for image collection. However, several instruments, such as BioRAD's direct viewing slit scanner, and Noran's AOD real-time scanner have also been used directly with NIH Image for data collection.
A set of macros has been provided for use with confocal images, as well as for the collection of fluorescent images using an integrating CCD. These are to be found in a separate file, entitled "Image Macros for CLSM". They can be used directly with NIH Image without modification. However, you may find that your personal preferences differ from mine, and you may modify the macros to better suit your needs. See Mark Vivino's excellent manual on Macro Programming with NIH Image , available from the usual FTP site ( zippy.nimh.nih.gov/pub/nih-image/ ).
If you wish these macros to load automatically each time you start NIH Image , place them in the same folder with the program or in the System Folder. Before you do that, Save the current version of "Image Macros" using an alternate name. Rename a copy of "Image Macros for CLSM" as "Image Macros".
The next time you start NIH Image, these macros will load as the default set. See the NIH Image manual for further details on loading macros.
IX. ADDENDA TO NIH IMAGE CONFOCAL IMAGING MANUAL
Addenda March, 1995
- Macros to merge sides of split screen BioRAD image. Macro to swap red and green slices of an RGB Stack.
- Transferring NIH Image RGB images to Adobe Photoshop 3.0
- Use of NIH Image for pre-evaluation of fluorescence--shutter and on-chip integration control. Provides a series of macros for controlling the shutter, on-chip integration, making new Stacks and related operations.
These macros are arranged in order of my personal preference (more or less). This order will probably not satisfy anyone else. If you rearrange the order of these macros, make sure that you are careful in regard to defining "Variables and Procedures". They must all be placed in the file in front of the macros that call them.
Define Procedures for Stacks
Some Universal Operations
macro '[P] Print Video'
macro '[Q] Get Path'
macro '[F2] Calibrate Image'
Operations on Confocal Files: Import, Split, Merge
macro [=] "Animate"
procedure ShowBioRadInfo(InfoOffset: integer)
macro '[F1] Import Biorad Z Series'
macro '[S] Merge Split BioRAD'
macro ' Color Merge Two Images'
macro ' Color Merge Two Stacks'
macro 'Separate SplitScreen Z Stack'
macro '[F5] RGB to Indexed '
macro '[W] Swap Red_Green'
macro ' Merge Two Stacks'
General Stack Operations
macro ' Make stack size front image'
macro '[F8] Flash to Stack
macro MakeStack w_Current Image'
macro 'Add Slice'
macro '[A] Add Black Slice'
macro '[D] Delete Slice'
Shutter Controls and On-Chip Integration
macro ' Autoshutter'
macro '[O] Open Shutter'
macro '[C] Close Shutter'
macro '[T] Trigger Shutter'
macro '[R] Reset Shutter'
macro '[G] Open Shutter-Grab-Close'
procedure Integrate (mode:string)
macro '[F6] Integrate On-chip Using Cohu'
macro '[F7] Integrate One Image on Cohu'
macro ' SetIntegrate:2 Frames'
macro '[F] Show nFrames'
macro 'Integrate White Light'
macro ' Live'
macro ' Average'
macro '[F1] Import Biorad Z Series'
macro '[F2] Calibrate Image'
Stack Modifications, Z-Projections, Stereo Pairs
macro ' Smooth Stack'
macro ' Sharpen Stack'
macro '[I] Invert Stack'
macro ' Reduce Noise Stack'
macro '[L] Apply LUT to Stack '
macro '[F0] Remove 0 and 255 from Stack'
macro ' Flip Stack Vertical'
macro ' Flip Stack Horizontal'
macro ' Clear Outside Stack'
procedure CropAndScale(fast:boolean angle:real)
macro '[E] Crop and Scale-Fast '
macro ' Crop and Scale-Smooth '
macro ' Rotate Left'
macro ' Rotate Right'
macro ' Rotate '
macro '[H] Reslice Horizontally'
macro '[V] Reslice Vertically'
macro '[F12] Z Projection'
macro '[F14] Make Stereo Views'
macro ' Invert LUT'
macro ' Log Tranform'
macro '[M] Gamma Tranform '
macro ' Reset LUT'
macro ' Set Pixels Red '
macro ' Nearly Gray LUT '
macro '[U] Move Slice Up'
macro '[Y] Move Slice Down '