Cell Imaging Facility

The LSCM uses a point light source (laser) and a pinhole to reject out of focus light. It enables optical sectioning of fluorescently stained samples to provide a 3-Dimensional view of the structure. For more information on confocal microscopy, please go to resource section.

Applications

1. To examine fluorescently labeled specimens for single or multiple colors localization of molecules within cell/tissue.
2. To carry out physiological studies such as ratio imaging, time-lapse recording of live cell/tissue.
3. Photo manipulation of the fluorescence signal (e.g. FRAP (photo bleaching after fluorescence recovering); FLIP (Fluorescence Loss Imaging after photobleaching), photoactivation/switching, etc.) to study the kinetics of molecules in living cells.

This system is mainly used for detection of fluorescence labeling in fixed samples. However, it is also equipped with necessary accessories (temperature, CO₂ controls, O₂) for live cell imaging. Additionally, the system is equipped with an FCS module which also allows imaging using the sensitive Avalance Photodiode Detector (APD, Zeiss Confocor 3, see FCS section below).

The instrument is equipped with a spectral detector which uses a PMT (Photon Multiplier Tube) array and optical grating elements to collect spectral information for fluorophores. Spectral information from 405nm to 720nm with step size of 10.7nm can be detected with the setup. Then, using the built-in, non-linear un-mixing tool, signals from different fluorophores can be separated mathematically. The system eliminates the need for emission filter and offers the advantage of being able to separate fluorophores with extensive emission spectra overlap. For example, GFP and YFP can be separated using this setup (which is impossible with conventional filter based imaging system). The system is also particularly useful for imaging highly auto-fluorescence sample with fluorescence imaging as auto-fluorescence has very different spectral signature than fluorophores.

Zeiss LSM 710 Specifications

This is a complex system with multiple capabilities:

1. Super resolution microscope with STED (STimulated Emission depletion) which is capable of obtaining fluorescence images with resolution down to <50nm.
2. Multiphoton microscope with tunable laser from 680-1064nm.
3. Fluorescence Lifetime Imaging system based on time domain for FLIM measurements.
4. Fluorescence correlative Spectroscopy system for detecting molecular associations.
5. Combination of optics and software module capable of obtaining fluorescence images down to ~120nm.

STED uses a donut shaped depletion laser to modify the point spread function of the beam down to sub-diffraction limit level to achieve super resolution microscopy at resolution below 50nm (STED principle). The system is equipped with 594nm, 660nm and 775nm depletion lasers for variety of dyes. For detailed system information, please visit the Leica website

Multi-photon microscope uses an ultra-fast pulsing laser to excite fluorescent molecules. Briefly, 2 (multi)-photon microscopy is based on the principle that a given fluorescent molecules which normally would only be excited by absorbing the energy of a single photon of certain wavelength can also be excited by combination of the energies produced by simultaneous absorption of 2 (or multi-) lower-energy photons of double (multiple) wavelength. The emission of the fluorescence is quadratically dependent on the excitation intensity. This steep dependence of absorption rate on photon concentration gives multi-photon Laser Scanning microscope the intrinsic three-dimensional resolution with the depth of field defined by:

1. intensity of the excitation light
2. numerical aperture of the lens used
3. the wavelengths of the lights.

This z-resolution is comparable to conventional Laser Scanning confocal microscope.

Advantages over confocal microscope

1. Longer wavelength used for excitation allows imaging deeper into thick specimens due to less scattering of longer wavelength (IR) light.
2. Photo-bleaching and damaging are confined to the focus points only.
3. It is possible to image UV dyes with infra-red light. This is particularly useful for live cell imaging where UV light is highly damaging to the cells

Therefore, multi-photon microscopy is highly suitable for imaging of live, thick specimens.
The system is also capable of measuring fluorescence life time with the equipped pulse lasers. The system is equipped with a white laser which is pulsed and could be tuned continuously from 470-690nm in addition to the IR laser with tuning range of 680-1024nm. In addition, the system is able to perform Fluorescence correlative spectroscopy, deconvolution and high speed imaging.

In the below image, COS cells were stained with an anti-nuclear pore protein conjugated to Star635P dye and imaged with Confocal (top) and STED (bottom) at the same optical plane to demonstrate the improvement in resolution with STED (scale bar 2um).

Leica Falcon SP8 STED system with Spectra-physics femto-second laser Specifications

The PerkinElmer UltraVIEW system (PerkinElmer Life Sciences Inc., MA, USA) is a Yokogawa (Yokogawa Corp. Japan) Nipkow spinning disk confocal system. It uses a spinning disk with multiple pinholes to achieve confocality (e.g. the rejection of out-of-focus lights). Nipkow disk refers to scanning disk with symmetrically placed spirals of pinholes through which illumination light is passed. Such pinholes split illumination lights into multiple 'minibeams'. When the disk spins, the light scans the sample in a raster pattern. Emission lights from the sample are detected through the pinhole to generate a confocal image of the sample that can be detected (with an EMCCD (Electron Multiplification Charge-Coupled Device) camera). Because the pinholes on a Nipkow disk must be placed up to 10 diameters apart in order to avoid cross-talking problem, the light throughput of traditional Nipkow disks is only ~1% of the light shining onto the disk. The Yokogawa scanhead has overcome this problem by using an innovative, collector disk containing microlenses placed in front of the Nipkow disk. The microlenses ensure that most of the light illuminating the disk is focused onto the pinholes. Transmission efficiency is thus increased from ~1% to 70% of the light falling on the disks allowing the sample to be illuminated with a sufficient quantity of light.

Advantages of the UltraVIEW system

The system has mainly 2 advantages over a conventional Laser Scanning confocal microscope:

1. Higher image speed
2. Lower photo-toxicity

The Laser Scanning Confocal Microscopy (LSCM) is a sequential scanning system where a single point of the specimen is illuminated at a time and LSCM uses a point detector (Photon Multiplier Tube (PMT)). Therefore, for LSCM, an expensive scanner is needed and electronic processing is necessary for image formation. The LSCM system is consequently relatively slow (typically 0.5-10sec/frame, slower scan can take up to a minute/frame). Whereas Nipkow disk system uses multi-point scanning disk and a cooled CCD camera which is a parallel array detector. This enables the Nipkow system to acquire images at higher speed (up to 360frames/sec) than with LSCM (typically 0.5-1 frame/sec). This will allow many rapid cellular processes (e.g. calcium imaging, vesicular trafficking) to be monitored in real time. More importantly, it has been demonstrated that the spinning disk confocal system reduces phototoxicity and photobleaching by as much as 5 folds comparing to a LSCM. It is speculated that the reduction in phototoxicity and photobleaching is probably due to the fact that the system splits the laser light into thousands of minibeams.

While the mechanism of such reduction in phototoxicity and photobleaching is still a subject of studying, spinning disk confocal is becoming increasingly the instrument of choice for live cell imaging. The system is particularly powerful for applications such as real-time (4-D) confocal microscopy, calcium signaling, vesicular trafficking; green fluorescence protein studies.

System Description

The system is based on a Zeiss Axiovert 200M inverted microscope with the following laser lines for excitation of fluorophores: 404nm, 440nm, 488, 514nm, 561nm and 633nm. It is equipped with a piezo focusing motor and all necessary accessories (Temperature, CO₂) for high speed live cell imaging. The scope is mounted with a high precision motorized stage for monitoring of multiple positions in a single sample holder. This system was upgraded with a FRAP (Fluorescence Recovery After Photobleaching) module in 2009.

Applications

The system is well suited for high-speed, live cell imaging with reduced phototoxicity. The system is particularly useful for 3-D timelapse to monitor fast cellular events because of the high focusing/acquisition speed than other systems in the Facility. With its FRAP module, FLIP, FRAP and photoactivation could be performed on the system as well.

Perkin Elmer Ultraview ERS FRAP Specifications

DIMs offer alternative ways of imaging biological samples with advantages of low photo-bleaching, high-photon efficiency and relatively low cost. In combination with deconvolution, high resolution imaging could be performed on these systems.

There are 6 DIM setups in the Facility. Each differs from the others by the software and hardware configuration for its main purposes:

• DIM station 1-4 contains Metamorph (Universal Imaging Corp , Molecular Devices), a sensitive, cooled CCD camera (Sensicam from Optikon , Cascade II EMCCD or CoolSnapHQ (Photometrics)) and a Zeiss microscope (AxioImagerZ, Axioplan IIM or Axiovert 200M). All of them are equipped with DIC (differential Interference Contrast) optics and fluorescence optics. The setups are mainly set for low-level fluorescent video microscopy. All these digital microscopes can perform multiple wavelength detection, 3-dimensional image acquisition and 3-D time lapse experiments. One setup is equipment with necessary filter wheels for Ca/pH ratiometric imaging and for FRET analysis with CFP and YFP dye pairs. It is also equipped with a live cell environment control that long term (days) timelapse can be performed on.

• DIM station 5 is composed of an Axioscope 2 from Zeiss for routine fluorescent examination and a high resolution digital color camera (Axiocam HR, Zeiss) for color digital photography.

• DIM station 6 is composed of a Zeiss Axiovert 100M, cooled CCD camera (Sensicam HE) and Metamorph software. The main difference between this setup and others is that this one is equipped with an Eppendorff microinjection system which allows injecting small amount of molecules in adherent cells. The system is, also available for routine fluorescence imaging.

• DIM station 7 contains an inverted fluorescence microscope (Olympus IX70) and a Zeiss Axiocam for routine sample examination as well as documentation. It could also perform time-lapse imaging with bright field (with phase contrast optics) if needed.

System Specifications

All of these scopes (except 7) have interchangeable optics and are based on either upright or inverted configurations. Please contact staff for your particular application needs.

This is a PALM CombiSystem mounted on a Zeiss Axiovert 200M microscope. The setup is used primarily for manipulating cells with laser or selectively picking up cells or sub-cellular structures for biochemical analysis (e.g. single cell PCR). The workstation consists of a pulsed nitrogen laser, which allows precise cutting or micro-dissection. It is also equipped with an infrared laser for trapping and positioning cells. More information about the system can be found on the Zeiss website.

Applications

1. Select morphologically identified cells/fragment of cells in pathological sample to study the sample biochemically.

2. Using the laser tweezers to physically position 2 cells together to study cell-cell interactions.

3. Selective ablation of cells in living tissue or cell culture.

The Facility is equipped with a 200kv JEOL 2100 Transmission Electron Microscope (TEM). TEM use an electron beam as light source (Lab6 crystal in this case) and the beam goes through (transmission) the specimen and the image is projected to an imaging device (e.g. fluorescence screen, film or CCD camera). The TEM can obtain resolution much higher than a light microscopy could offer (sub nm resolution versus ~200nm) due to the much shorter wavelength of the electron beam than visible light.

1. Fixation of the specimen which preserve the sample as close to the native status as possible before viewing under the TEM. This are usually achieved through one of the following methods: i) Chemical fixation which typically use cross linker molecules such as glutaraldehyde to fix major cellular components. ii) Cryofixation which involved in freezing the sample so rapidly that the samples are frozen in vitreous state. Some of the fixation reagent also enhance the contrast of the specimen under the TEM (e.g. OsO₄).

2. Contrast enhancing: since most biological material yield low contrast under electron beam, it is necessary to apply some heavy metal reagents to increase the contrast of the sample. Some of the commonly used one are OsO₄ (for lipid staining), Uranyl acetate (for nucleus and negative staining); Lead citrate for general electron opaque contrast enhancing.

3. Resin infiltration and plastic embedding: Since typical cellular structure is too thick to observe under TEM, it must be sectioned to thin sections (typically 70nm) prior viewing under TEM. This requires the specimen to be embedded in plastic resin. Typically, the specimen goes through steps like dehydration, resin infiltration and polymerization of resin (either in an oven or under UV light).

4. Thin sectioning (ultra microtomy). Typical biological TEM has an acceleration voltage of 200kv or below and at this acceleration voltage, electrons can penetrate a specimen of thickness of less than ~200nm. A special instrument is used with a diamond knife to cut the specimen down to ~70-100nm for TEM observation and the sections are placed on to a specimen grid.

5. Post staining if needed. Additional contrast enhancement is possible at this stage.

6. Section stabilization: Many biological samples with resin are unstable under TEM beam (either due to local charging or heating generated by the electron beam). A thin layer of carbon (1-5nm) can be applied to the sections in order to increase the stability of the sections under the electron beam.

TEM images are formed by the interactions of electrons and the specimen and the TEM can operate in different modes to extract different information from the sample:

• Brightfield is the most common operation mode where most of the contrasts are formed by absorption of the electrons by the specimen yielding an image of different levels of shade which reflects the degree of electron absorptions of the sample.

• Darkfield mode explores the scattered electrons. A bright spot in the image means the structure is highly scattering (e.g. a gold particle).

• Electron Energy Loss Spectroscopy (EELS) detects the electrons which under inelastic interactions with the specimen (therefore loss some energy) and gates electrons of different energy level to the detector. Using this information, element composition of the specimen can be mapped. This mode is also very effective way to enhance contrast by filtering out the scattered electrons from the final image.

Applications

TEM can be used to obtain details view of a specimen at atomic resolution level. For biological samples, TEM can give unambiguous identification of cellular compartments and organelles. In combination with specific molecular marker (e.g. antibody) and electron dense labels (immunogold particles or Nanoparticles), it is possible to identify association of a particular molecules with a specific organelle or with another type of molecules. With Tomography, 3-dimensional information can be obtained. With EELS, an elementary composition of the specimen can be mapped. For biological samples, for example, phosphorous rich and nitrogen rich domains can be mapped. The TEM is also equipped with cryo sample holder to observe sample under cryo-conditions.

System Specifications

• Cryo sample holder for investigating sample at liquid nitrogen temperature.
• Double tilt sample holder for Tomography.
• Gatan GIF Tridieum filter for EELS for element mapping.
• High Angle Annular Dark field detector for dark field imaging.
• Necessary TEM sample preparation for biological samples. This includes:
  • High pressure freezer for cryofixation.
  • Freeze substitution unit for substituting water from biological sample under cryo condition.
  • Ultramicrotome with Cryo attachment for ultramicrotomy either at room temperature of low temperature
  • Immunogold labeler to perform immunogold labeling.
  • Microwave automatic tissue processor for rapid tissue processing for TEM.
  • Carbon coater with glowing discharge and metal shadowing capability.
  • The scope is equipped with Gatan DigitalMicrograph software with Low-dose function for radiation sensitive sample (cryo specimen), Montaging for reconstruction of large field of view. For tomography, it uses SerialEM.

Fluorescence is a cyclical process where lifetime of the fluorophore is not only the property of molecule itself but also a function of its environment. In situation such as Fluorescence Resonance Energy Transferring (FRET), the lifetime of donor molecule will be shortened. Therefore, FLIM can be used as a way to measure molecular interactions. Unlike conventional FRET analysis, FLIM measurement is not subjected to fluorescence photobleaching or concentration variations which are difficult to control in live cells.

The FLIM module is an attachment to the Zeiss NLO 510 system which uses time-correlated single photon counting technique to construct the decay curves of fluorophore in every pixel of the image. The system is equipped with a Hamamatsu RS-39 Multi-channel plate detector, a filter wheel and a SPC730 photon-counting board from Becker Hickl for photon counting. The system uses a photodiode to obtain synchronization information from the laser pulses to construct the fluorescence decay curve.

Applications

The main application for the setup in biology is for FRET analysis where donor life time can be measured both in presence and absence of acceptor molecules. Then the FRET transfer efficiency can be calculated according to the following formula: - Et =1- t _D,A/ t_D

Where t _D,A is the life time of donor molecule in presence of acceptor and t _D is the life time of the donor molecules in absence of acceptor.

The system is integrated part of multi-photon microscope. There is no special need for specimen preparation other than that the specimen must be suitable for confocal observation. As all FRET analysis, all control samples are needed (e.g. donor alone, acceptor alone, donor acceptor combined and specimen without any staining for auto-fluorescence).

The system does not have an overly user friendly interface, therfore it is only available by assisted use. Please contact staff for more details.

FCS is a spectroscopic technique for the study of molecular interactions in solution. FCS monitors the random motion of fluorescently labelled molecules inside a defined volume element irradiated by a focused laser beam. These fluctuations provide information on the rate of diffusion (diffusion time) of a particle and this, in turn, is directly dependent on the particle's mass. As a consequence, any increase in the mass of biomolecules, e.g. as a result of an interaction with a second molecule, is readily detected as an increase in the particle's diffusion time.

Applications

Because FCS measurement is made through diffraction limited volume, FCS can be used to study molecule-molecule interactions in living cell. The system can be used to study:

1. Receptor-ligand interactions

2. Transcription factor-DNA interactions

3. Lipid-protein interactions, etc.

The FCS system at the Facility is an integrated part of the Multi-photon and the LSM710/Zeiss LSM NLO systems with all the laser lines for common fluorophores and 2 detectors which enable cross correlation analysis.

Micro-injection is one of the techniques to introduce minute amount of substance into live cell/tissue. The Facility is equipped with an Eppondorff micro-injection system mounted on a Zeiss Axiovert 100M fluorescent microscope. It is suitable to inject small amount of substance (e.g., antibody, dye, drug, etc.) into adherent cultured cells. The system is also equipped with a cooled CCD camera (Sensicam) and Metamorph for fluorescence imaging.

The Facility is equipped with various up-to-date computer workstations for image processing. These include several high end workstations with powerful graphics and large amount of RAM and hard drive space dedicated for image processing and analysis. Here is a list of software tools available for the Facility users:

Deconvolution software

• Autodeblurr from Media Cybergenetics
• Huygens essential from SVI
• SoftWoRx from API

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3-D image processing and analysis software

• Imaris from Bitplane
• Volocity from PerkinElmer

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2-D image processing and analysis and image acquisition software

• Metamorph from Molecular devices
• ImageJ
• Adobe Photoshop
• Zeiss Axiovision

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Some of the computers are equipped with Zeiss LSM software (ZEN) for offline processing of confocal/FCS data.