The Hospital for Sick Children
The Imaging Facility

ION MEASUREMENTS

QUESTION: How can I measure the pH of a cellular compartment?

ANSWER: pH sensitive fluorescence proteins (e.g. eGFP, pHluorin) or dyes (e.g. SNARF, Oregon Green) can be targeted to specific organelles (e.g. lysosomes, the Golgi complex or recycling endosomes) and used to measure pH in situ

PRINCIPLE: The spectral characteristics of some fluorophores change in a predictable manner according to their local pH environment. By comparing the excitation or emission intensities of a pH-sensitive fluorophore in a given compartment to a standard calibration curve, the local pH can be readily determined. In addition, some fluorophores demonstrate a pH-dependent change in the lifetime of their excited state. By comparing the fluorophore lifetime in a given compartment to a standard calibration curve, the local pH can be determined.

EQUIPMENT: Zeiss epifluorescence microscope, Lambert FLIM

 

QUESTION: How can I measure calcium/magnesium or other ionic concentrations in single cells and/or organelles?

ANSWER: Calcium/magnesium-sensitive fluorophores (Fura, Indo, etc.) as well as other ion-sensitive probes can be used in live cells to measure ion concentrations.

PRINCIPLE: Certain fluorescent dyes selectively bind calcium, magnesium, sodium or other ions, resulting in a concentration-dependent change in their spectral properties. These dyes can be used in live cells to measure ionic concentrations in specific compartments by comparing their in situ intensity to a standard calibration curve. In addition, some fluorophores demonstrate an ion-dependent change in the lifetime of their excited state. By comparing the fluorophore lifetime in a given compartment to a standard calibration curve, the local ion concentration can be determined.

EQUIPMENT: Zeiss epifluorescence microscope, Lambert FLIM

 

QUESTION: How can I detect calcium sparks in live cells?

ANSWER: Highly sensitive alcium-specific probes (e.g. Fluo-3 and Rhod-2) can be used in live cells to detect rapid changes in calcium concentration. 

PRINCIPLE: Upon binding calcium, selective probes like Fluo-3 and Rhod-2 (that have a large dynamic range) undergo a concentration-dependent change in their fluorescence intensity. In combination with confocal scanning, calcium sparks can be detected in the millisecond time range.

EQUIPMENT: Nikon A1R confocal microscope, Zeiss LSM510 confocal microscope, Zeiss epifluorescence microscope

 

QUESTION: How can I measure changes in membrane potential?

ANSWER: A variety of fast- and slow-responding electrical potential-sensitive fluorescent dyes (e.g. oxonols, cyanines) can be used to measure membrane hyperpolarization and depolarization. 

PRINCIPLE: Certain fluorescent dyes display changes in their intensity as a function of the membrane potential. Such dyes can be used in combination with other, organelle-targeted fluorophores to report changes in membrane potential using FRET. In addition, some fluorescent proteins have been engineered to report the transmembrane potential.

EQUIPMENT: Nikon A1R confocal microscope, Quorum spinning disk confocal microscope, Zeiss epifluorescence microscope

 

QUESTION: How can I measure the concentration of my protein in live cells?

ANSWER: Fluorescence correlation spectroscopy (FCS) can be used to measure protein concentrations as low as the sub-nanomolar range.

PRINCIPLE: In FCS, fluorophores diffusing in and out of a known confocal volume are observed. By temporally auto-correlating the intensity and duration of the fluorescence fluctuations within the confocal volume, the concentration of a given fluorophore can be calculated.

EQUIPMENT: Nikon A1R confocal microscope

 

 

PROTEIN MOBILITY/TRAFFICKING

 

QUESTION: How can I track a specific subpopulation of cells over time?

ANSWER: Cells transfected with photo-activatable GFP (or equivalent variants such as the Eos family of proteins) can be ‘turned on’ selectively and tracked over time.

PRINCIPLE: Photo-activatable GFP normally exists in the dark state, becoming fluorescent only upon excitation with UV light. Localized illumination of specific populations of cells with UV light allows for visualization of only those cells, allowing for very precise tracking of cell fate.

EQUIPMENT: Nikon A1R confocal microscope, Quorum spinning disk confocal microscope

 

QUESTION: How can I measure protein mobility?

ANSWER: Fluorescence recovery after photobleaching (FRAP) experiments can be used to measure the diffusion coefficient of slow moving proteins (e.g. membrane proteins). For fast-moving proteins (e.g. cytosolic proteins), FCS microscopy can be used instead. Direct measurements of protein mobility can also be made using single-molecule imaging and tracking.

PRINCIPLE: In FRAP microscopy, a specific population of fluorophores are selectively photo-bleached by a fast, intense pulse of excitation light. The reappearance of fluorescence within this area is monitored as bleached fluorophores diffuse out and neighbouring (unbleached and therefore fluorescent) molecules diffuse in from the surrounding regions. By analyzing of the rate and extent of recovery of fluorescence in the bleached region, diffusion coefficients can be calculated and the fraction of mobile molecules determined.

In FCS experiments, fluorophores diffusing in and out of a known confocal volume are observed, and various parameters of protein mobility can be calculated, based on their transit time through the observation volume. Specialized software can also analyze time-lapse microscopy images to track single molecules over time, providing direct estimations of diffusion coefficients and patterns (e.g. Brownian motion vs. confinement or tethering).

EQUIPMENT: Nikon A1R confocal microscope, Quorum spinning disk confocal microscope, Zeiss epifluorescence microscope

  

QUESTION: How can I visualize vesicle exocytosis?

ANSWER: Because total internal reflection fluorescence microscopy (TIR-FM) microscopy is limited to events occurring within 200 nm of the coverslip, it is an ideal tool for imaging exocytic events at high resolution.

PRINCIPLE: When incident light is directed towards the coverslip at a certain angle, the laser beam is reflected and only an evanescent field, extending nearly 200 nm from the coverslip, is produced. Therefore, only fluorophores near the glass surface are excited using this technique. By loading exocytic organelles with pH-sensitive dyes (e.g. acridine orange) or with other probes, exocytosis occurring at the ventral surface of cells can be visualized using TIRF microscopy.

EQUIPMENT: Olympus TIRF microscope

 

QUESTION: How can I study protein trafficking using fluorescence microscopy?

ANSWER: Confocal microscopy can be used to image fluorescently labeled proteins within the cell. Observing the localization of your protein of interest at various time points, in combination with fluorescence labeling of known cellular compartments will provide insight to protein trafficking.

PRINCIPLE: Fluorescently labeled proteins that are known to reside in specific cellular compartments can be used as markers for those organelles. By labeling your protein of interest with a different fluorophore, along with known organelle markers, protein trafficking pathways can be elucidated at various time intervals. The dynamics of protein or organellar trafficking can be examined by live cell imaging for periods ranging from several seconds to several hours.

EQUIPMENT: Spinning disk confocal microscope, Nikon A1R confocal microscope, Olympus TIRF microscope

 


MOLECULAR/CELLULAR INTERACTIONS 

 

QUESTION: How can I determine if two endogenous proteins are part of the same complex/organelle?

ANSWER: Confocal microscopy can be used to determine whether two fluorescently labeled proteins co-localize.

PRINCIPLE: Proteins that are fluorescently labeled using secondary antibodies of different emission wavelengths (e.g. green and red) can be imaged simultaneously on a confocal microscope. Overlap of the two fluorophores in a composite image suggests that this pair of proteins reside within the same complex (defined by the resolution limits of optical microscopy method employed). Analysis of these images with specialized software can quantitate the extent of colocalization observed.

EQUIPMENT: Spinning disk confocal microscope, Nikon A1R confocal microscope, Olympus TIRF microscope, Volocity software

 

QUESTION: How can I determine if two proteins interact?

ANSWER: FRET microscopy can reveal if two proteins are within 10 nanometres of each other, and therefore likely to be directly interacting.

PRINCIPLE: Fluorescence resonance energy transfer (FRET) is a process in which  transfer of energy occurs from an excited fluorophore (donor) to a second fluorophore (acceptor) that is in close proximity (less than 10 nm). This energy transfer can be detected in a variety of ways. Sensitized emission FRET is based on the detection of the increased emission of the acceptor fluorophore resulting from transfer of energy from the excited donor. Acceptor photobleaching FRET measures increases in the fluorescence intensity of the donor molecule following photobleaching of the acceptor.

Fluorescence lifetime imaging (FLIM) examines changes in the donor fluorescence lifetime caused by energy transfer. FLIM measurements of energy transfer are generally considered the most practical and reliable method for FRET.

EQUIPMENT: Lambert FLIM microscope, Nikon A1R confocal microscope, Zeiss epifluorescence microscope.

 

QUESTION: How can neuronal circuits be imaged?

ANSWER: The Brainbow technique allows individual neurons to be labeled in multiple, distinct colours. Using confocal microscopy, three-dimensional reconstructions of multi-coloured neural circuits can be generated.

PRINCIPLE: The Cre/loxP recombination system can be used in conjunction with genes encoding different fluorescent proteins to generate stochastic combinations of 2 or more fluorescent proteins inside individual neurons. This technique can generate more than 100 unique colours. Imaging these samples using a confocal microscope with spectral unmixing allows for complete neural circuits to be traced in various regions of the brain.

EQUIPMENT: Nikon A1R confocal microscope

 

 

IMAGE ANALYSIS 

 

QUESTION: How can I determine differences in protein expression/localization?

ANSWER: Image analysis software (e.g.  Volocity) can be used to measure pixel intensity values.

PRINCIPLE: Digital images are constructed from a matrix of tiny light sensitive elements called pixels. The intensity of any given pixel is directly proportional to the number of incident photons. Thus, regions of high pixel intensity reflect a higher abundance of fluorophores (i.e.molecules) compared to regions of low pixel intensity. Images acquired can be analyzed in an automated fashion to measure pixel intensities (i.e. expression/labeling) across the entire image or in specific regions of interest, for multiple fluorophores.

EQUIPMENT: Perkin Elmer Volocity software

 

QUESTION: How can I count cells in my images?

ANSWER: Image analysis software can be used to generate cell counts for both bright-field and fluorescence images.

PRINCIPLE: For bright-field images, colour deconvolution is used to analyze staining intensity only on the chromogen channel. Based on these data, nuclei can be classified into negative, weak positive, medium positive and strong positive categories. For fluorescence images, cells can be counted based on intensity thresholding, size filtering and cluster splitting algorithms. This analysis can be done both in 2D and 3D.

EQUIPMENT: 3D Histech Pannoramic Viewer, Perkin Elmer Volocity

 

QUESTION: How can I improve the resolution of my fluorescence images?

ANSWER: Confocal and wide-field fluorescence images can be mathematically deconvolved to remove out of focus blur, thereby improving resolution.

PRINCIPLE: Deconvolution is a computational image-restoration technique that can remove out of focus blur to improve contrast and resolution. Using a mathematical description of how light spreads around a point source of light (termed the point spread function), out of focus light can either be removed or reassigned to its origin, resulting in improved lateral and axial resolution. Deconvolution can be applied to virtually any sort of fluorescence images.

EQUIPMENT: Perkin Elmer Volocity


OTHER IMAGE ANALYSIS/EDITING TOOLS AVAILABLE:

Quantitation: Perkin Elmer Volocity, Bitplane Imaris

Colocalization: Perkin Elmer Volocity, Bitplane Imaris

3D Rendering: Perkin Elmer Volocity, Bitplane Imaris

Image Stitching: Perkin Elmer Volocity

Particle Tracking: Mathworks MatLab

Time-Lapse Movies: Perkin Elmer Volocity, Bitplane Imaris

Manuscript/Presentation Preparation: Microsoft Office, Adobe Creative Suite

Custom Applications: Mathworks MatLab

 

 

BIOCHEMICAL/POPULATION ASSAYS 


QUESTION: How can I test for cell viability?

ANSWER: The MTT assay is one example of a colorimetric assay for assessing cell viability. Fluorescence based assays can also be used. Both approaches can be performed using microplate readers, enabling analysis of large populations of cells.

PRINCIPLE: MTT is a pale yellow substrate that is cleaved by living cells to yield a dark blue formazan product. This process requires active mitochondria and is therefore a popular method to assess proliferation or cytotoxicity. Microplate readers can be used to measure light absorbance across many samples of a multi-well plate, in both endpoint and kinetic modes. The measured output, when coupled with a standard curve, can yield quantitative information on cell viability. Analogous assays using fluorescence reporters can also be used.

EQUIPMENT: Molecular Devices SpectraMax Gemini EM/Versamax 190 Plate Readers

 

 

QUESTION: How can I measure gene expression?

ANSWER: The luciferase reporter assay can be used to study gene expression rapidly and with high enough sensitivity to quantify even small changes in transcription. This assay can be performed using a luminometer.

PRINCIPLE: Luciferase is an oxidative enzyme that catalyzes the conversion of luciferin to oxyluciferin. A byproduct of this reaction is the rapid release of energy in the form of light. By cloning the regulatory region of your gene of interest upstream from the luciferase gene in an expression vector, expressing it in cells, and performing the assay, gene expression can be measured as a function of light output. A luminometer can measure the light produced in a quantitative and highly sensitive manner.

EQUIPMENT: Molecular Devices SpectraMax L

 

QUESTION: What is the best way to quantitate Western blots?

ANSWER: Camera-based imaging of Western blots provides a highly quantitative, highly sensitive method of analyzing chemiluminescent or fluorescently labeled Western blots.

PRINCIPLE: Photons generated from chemiluminescent or fluorescently labeled Western blots are collected on highly sensitive digital detectors. The accuracy, linearity and wide dynamic range of these detectors allows for both strong and weak bands to be detected on the same blot and quantitated precisely in a linear fashion, without the use of film. An added level of sensitivity can be also be achieved by using infrared fluorophores, enabling the detection of protein levels as low as one picogram.

EQUIPMENT: Li-Cor Odyssey FC, Molecular Dynamics Typhoon Phosphorimager

  

QUESTION: How can I excise specific populations of cells from tissue for further analysis (e.g. PCR/RT-PCR)?

ANSWER: Laser microdissection microscopy can be used to selectively dissect specific populations of cells from tissues. Isolated cells can be used for a variety of subsequent experiments, including RT-PCR or qPCR.

PRINCIPLE: Tissue sections labeled via immunofluorescence or immunohistochemistry mounted on specialized slides can be microdissected using a high-powered ultraviolet laser to isolate specific populations of cells. The dissected population is then transferred/captured to specialized tubes for further analysis

EQUIPMENT: MMI Laser Microdissection microscope