Confocal microscopy

 

Why use a confocal microscope?

 

Confocal microscopy offers several advantages over conventional optical microscopy including;

 

i.                    Increased effective resolution: Confocal microscopy produces images of improved resolution, up to 1.4 times greater than standard microscopy and also has a higher level of sensitivity compared to conventional microscopes, due to highly sensitive light detectors and the ability to accumulate images captured over time.

 

ii.                  Clear examination of thick specimens (Controllable depth of field): Double and triple labels can be collected with a confocal microscope. Since these images are collected from an optical plane within the sample, precise colocalizations can be performed.

 

iii.                  Reduced blurring of image from light scattering: Eliminates the out of focus "haze" normally seen with a fluorescent sample. Fine detail is often obscured by the haze and cannot be detected in a non-confocal, fluorescent microscope.

 

iv.                 Improved signal to noise ratio

 

v.                   Z-axis scanning: The confocal microscope has a stepper motor attached to the fine focus, enabling the collection of a series of images through a three dimensional object. These images can then be used for a two or three dimensional reconstruction.

 

vi.                 Magnification can be adjusted electronically

 

 

Purpose (Applications):

 

Confocal microscopy is used in a large variety of scientific fields.

 

  1. Live cell imaging. This is often done by making use of a green fluorescent protein (GFP) or other fluorochromes attached to cellular components (proteins, genes, cell structures) and imagining their position or movements.

 

  1. Study of cellular/subcellular functions:

 

i.                     Esterase activity

ii.                   Oxidation reaction

iii.                  Intracellular pH

iv.                 Intracellular Calcium

v.                   Phagocytosis and Internalization

vi.                 Apoptosis

vii.                Membrane potential

viii.              Cell-cell communication (Gap junction)

 

  1. Study of conjugated antibody (colocalization study)
  2. DNA/RNA
  3. Organelle structure
  4. Cytochemical identification
  5. Probe rationing

 

 

How does a confocal microscope work?

 

Confocal microscopy generally uses a monochromatic light source (usually a laser) to excite fluorescent dyes. The light source is then focused into a single plane or spot which can be scanned across the specimen on one focal plane at a time. The thickness of the spot or plane of light is usually between 0.25 and 2 mm. The light is then scanned across the specimen by an oscillating mirror. In essence the confocal microscope optically sections the specimen. The specimen can then be raised and lowered so that the light can slice through at different planes. This process eliminates the problem of out of focus fluorescence. Subsequently, images from different focal planes can be digitized and overlayed to produce a single composite image with superior contrast or the images can be stacked and stored digitally as a three dimensional image.

 

 

 

 

A laser is used to provide the excitation light (in order to get very high intensities). The laser light (blue) reflects off a dichroic mirror. From there, the laser hits two mirrors which are mounted on motors; these mirrors scan the laser across the sample. Dye in the sample fluoresces, and the emitted light (green) gets descanned by the same mirrors that are used to scan the excitation light (blue) from the laser. The emitted light passes through the dichroic and is focused onto the pinhole. The light that passes through the pinhole is measured by a detector, i.e., a photomultiplier tube (PMT).

 

 

Glossary and techniques:

Laser Systems: The lasers commonly employed in laser scanning confocal microscopy are high-intensity monochromatic light sources.

Pinhole: In confocal microscopy, variable diaphragm pinhole apertures placed near the light source and detector enable the microscope to produce thin optical sections of focal planes in the specimen. A pinhole refers to a small break in the coating (primarily arising in the thin-films of interference filters) produced by dust particles or debris on the substrate during application. Pinhole size is measured against a standard under specific conditions using a high-intensity illumination.

Photomultiplier tube (PMT): An electrical device designed to collect and amplify photon signals. Incoming photons strike a target in the face of the photomultiplier to liberate free electrons, which are accelerated onto a dynode that in turn liberates an amplified stream of electrons. Several dynodes are arranged in series to produce a tremendous degree of amplification from each original photon and then transmit the signal to a processing circuit. Unlike area-array detectors such as charge-coupled devices (CCDs), photomultipliers do not form an image.

Bleed-through (Cross-over): Bleed-through of fluorescence emission, due to the broad spectral profiles exhibited by common fluorophores, is a fundamental problem that must be addressed in laser scanning confocal fluorescence microscopy. The phenomenon is most often manifested by the emission of one fluorophore being detected in the channel or through the filter combination reserved for a second fluorophore.

Bleed-through occurs when unwanted wavelengths are transmitted through an optical filter designed to block them. The effect usually occurs when the design of a filter does not allow complete destructive interference of wavelengths excluded from the bandpass region. Bleed-through is also a problem when the incident light angle is highly oblique with respect to the filter surface, or when the excitation and emission spectra of fluorescent dyes overlap.

z-series and 3D image:

 

A popular mode of optical microscopy in which a focused laser beam is scanned laterally along the x and y axes of a specimen in a raster pattern. The emitted fluorescence (reflected light signal) is sensed by a photomultiplier tube (PMT) and displayed in pixels on a computer monitor. The pixel display dimensions are determined by the sampling rate of the electronics and the dimensions of the raster. Signal photons that are emitted away from the focal plane are blocked by a pinhole aperture located in a plane confocal with the specimen. This technique enables the specimen to be optically sectioned along the z axis.

 

A 3D image created by adding together multiple optical sections acquired along the z-axis of a confocal microscope. In conventional widefield fluorescence microscopy, images of three-dimensional specimens are often blurred due to emission occurring away from the immediate focal plane. The same images are often remarkably sharp when gathered with a confocal microscope.

 

Scanning modes:

Among the common scanning modes are point, line, free line, parallel plane, and rectangle scanning over one or more dimensions. Line scanning along the x, y, or z axis, or diagonally between axes, provides intensity information across a single set of coordinates in the lateral or axial dimension. This scanning mode is useful for obtaining accurate quantitative information about rapid physiological events, such as calcium waves or sparks.

 

 

Sequential scan mode:

 

In sequential scan mode, images will be recorded in a sequential order instead of acquiring them simultaneously in different channels. Each sequence can be recorded using an individual set of user-defined parameters to optimize performance and image quality. e.g. simultaneous image acquisition of double or triple-stained samples can result in crosstalk since all dyes will be excited at the same time. Defining parameter sets specific for each dye and executing them in a sequential order can eliminate this crosstalk

 

 

Procedure (That I follow in my lab. for capturing confocal image)

 

I infect the mice [Wild type (C57BL/6) vs CXCL10 knock out mice] with 1000 pfu/ml HSV-1 (3 µl) per eye. At different times post infection at day 3, 5 and 7, I euthanize the mice and harvest the cornea for detecting infiltrating CD4 and CD8 T cells (and also for colocalization of CXCL10 and virus antigen) using Confocal microscopy.

 

A. Preparation of Corneal mounts:

 

Fixation:

 

  1. Harvest the cornea from uninfected and infected mice.
  2. Place each cornea in 4% paraformaldehyde at 4oC for 20 min.
  3. Wash 5X in 1X PBS.

 

Blocking:

 

  1. Block cornea with 100 µl PBS-BGEN with 1 µl FcBlock each for 2 hours.
  2. Add 5 µl Rat serum and incubate at least 5 hours upto overnight.
  3. Remove block.

 

Staining:

 

  1. Stain with 100 µl of 1:50 dilution of antibody (CD3-Alexa633, CD4-FITC, and CD8-TRITC) in PBS-BGEN overnight.
  2. Wash 5X at least 15 min. each with cornea wash buffer.
  3. Wash 1X with PBS.

 

Fixation:

 

  1. Fix with 1 % paraformaldehyde at 4o C for 20 min.
  2. Wash 5X with PBS.

 

DAPI:

 

  1. Add 1 drop of Vectashield with DAPI to each cornea in tube and soak overnight. Make sure that cornea is suspended in DAPI.
  2. Make star cut in cornea and mount with Vectashield.

 

 

 

B. Using confocal microscopy (FV500):

 

  1. Turn on the machine in following order;

 

    1. Xeon bulb power
    2. Laser box power
    3. Shutter (for choosing fluorescent dyes DAPI/TRITC)
    4. Stage (Xy) power
    5. Computer power
    6. Transmission light (Generally, we don’t use it)
    7. Laser (filter)- 405 for DAPI
    8. Laser (filter)- 568 for FITC
    9. Laser (filter)- 647 for Alexa-633

 

  1. Place your slide on stage (upside down) and select 10X or 20X objective to focus.
  2. After focusing at lower objective, select 40X or 60X objective to focus. While using 40X or 60X objective, use a drop of water on stage and place your slide upside down.

 

 

  1. Select “FluoView” software on computer.
  2. Select   “Acquisition Tab”

 

- Click on “Dye”: Select dyes (DAPI, FITC, TRITC or Alexa-633)

- Apply

- Click on Laser for chosen dyes: Apply

                        - Focus: X2, X4

                        - C.A. : don’t change C.A. (pinhole) normally

                        - Size: 512X512; high resolution 1024X1024

                        - Mode: Normal, Kalman

  (Generally, I use Kalman for sequential scan; select frame Kalman and   

   select 2 or 3)

-         Mode: Sequential scan 123

 

 

  1. Checking bleed through: (Do bleed through check before sequential scan)

 

Focus your slide and select only one laser at a time and detect the bleed through for that laser. If you detect bleed through, then follow the following instructions;

 

A. Select one laser, turn off others

     Press Focus, Press Stop

 

i. Select channel 1 (DAPI), Select DAPI laser

ii. Press coloring tool

 

iii. Select following intensity mapping;

Gamma 1.0

Press Hi-Low               High (red) ---------- Low (black or blue)

Adjust PMT, Gain and Offset

Press Focus, press Stop scan

 

 

B. Likewise select channel 2 and channel 3:

 

Click coloring tool (LUT), click Hi-Low and set up PMT, Gain, and Offset, select Gamma 1.0

 

C. After adjusting bleed-through, set coloring tool as according to dye color e.g. Blue for DAPI, Green for FITC and Red for TRITC. Apply.

 

7. Perform sequential scan selecting all dyes, lasers and selecting all required parameters.

8. After scanning, make 3D image using software. Analyze the observation.