Purpose:      To introduce  students to the light microscope as an observational and analytical tool in cell biology.

Introduction:

The light microscope is perhaps the single most important instrument used in cell biology.  It is used under bright field conditions to study the organization of cells in fixed and stained cells. This exercise involves the use of the light microscope under bright field conditions.  Cells will be fixed, stained and observed.

In addition to bright field mode, the inverted microscope we use in this lab also provides phase contrast conditions. Phase contrast microscopy converts phase shifts in light passing through a transparent specimen to brightness changes in the image. It enables the visualization of living cells, unstained cells and various cell organelles. The microscope in phase contrast conditions will be used to quantitatively determine the number of cells in a suspension using a special device known as a hemocytometer, or cell counting chamber.

Procedure:

A. The anatomy of the microscope

The light microscope consists of an optical system and a body.   The  body  is composed of a base which rests on the desk, a stage which supports the material to be viewed, and a tube through which the specimen is viewed.

The optical system begins with a light that shines through the condenser lenses on the top.  The function of the condenser is to focus the light source onto the specimen on the stage.  Light travels through the specimen to the objective. This lens magnifies the image of the specimen which is carried to the eyes by the oculars or eyepieces (Figure 1).

Figure 1.  Inverted light Microscope

B. How to focus

1.  With a piece of lens paper, clean the eyepieces and objectives.

2.  Slide the lower power objective into place.

3.  Place a specimen on the microscope stage.

4.  Watching from the side and being careful not to drive the objective into the coverslip, lower the objective as far as you can.

5.  Viewing through the eyepieces, focus up (raise the objective) until a semblance of the specimen is seen.

6.  Adjust the iris diaphragm, filter size or light control to optimize the lighting.

7.  With a bit of fine focusing, you are all set.

Some precautions in using the microscope

l One should take great care in handling the microscope.  When moving it from one place to another, it should be gripped tightly with both hands.

l If necessary, the lenses should be cleaned with lens paper (but not paper towel!).

l Always slide the lower objective into place and lower the stage, before inserting a microscope slide.

l As a general rule, you should start focusing with the 10X objective unless the large size of the object requires starting with the 4X objective.  Take great care in switching from a low-power objective to a high-power objective lest the lens hit the slide.

l Remember that the image is inverted. When you move the slide to the left, your image will move to the right.

l Your microscope is parcentric and parfocal.  This means that if an object is centered and in sharp focus with one objective, it will be centered and in focus when another objective is rotated into the viewing position (without adjusting the position of the stage),  e.g.  from  4X  to   10X  to  40X  to  increase  magnification.  However,  slight adjustmentstore-center andre-focus (with the fine-focusing knob) may be necessary.

NEVER use the coarse adjustment knob to focus the objective when you are using 40X or higher-power objectives.

l Total magnification = the power of the objective (4X, 10X, 40X…) multiply by the power of the eyepiece (usually 10X) , i.e. 40X, 100X and 400X.

l When putting the microscope away, adjust the stage to its lowest position, leave the

lowest objective in place, and return the microscope to the correct cabinet cubicle.

C. Measuring the field size by stage micrometer

It is possible to estimate the size of an object by comparing it with the diameter of the field of view.  To do this, it is first necessary to measure the size of the field.

1.  With the 40X objective engaged,  place stage micrometer (a short and transparent ruler) over the opening in the center of the stage so that the lines are visible through the microscope.

2.  Move the ruler so that a horizontal millimeter (mm) mark is just visible at the top edge of the circular field of view.

3.  Count the number of millimeters from the top to the bottom point.   If the top point of the field does not line up with one of the horizontal markings, estimate the fraction of a millimeter. This is the diameter of the 400X field of view.  Record your measurement in millimeters (mm) and in micrometers (μm) on the Datasheet (P.8) .  Note that the diameter of the field is less than 1 mm.

4.  Calculate the diameter of the 100X field by the following equation:

diameter of the high power field                                low power magnification

    =                                                        

diameter of the low power field                                 high power magnification

D. Calibration of ocular micrometer and measuring cell length or diameter

An ocular micrometer is a glass disk that fits in a microscope eyepiece that has a ruled scale, which is used to measure the size of magnified objects.   As the physical length of the marks on the scale varies with the degree of magnification, it requires calibration with stage micrometer.

1.  Place an ocular micrometer  in the eyepiece of the  microscope, according to the instructions of TAs or technician.

2.  Put a stage micrometer on the microscope stage.

3.  Focus on the scale divisions under low power.  Both scales should be sharply defined in the field of view.

4.  Turn the eyepiece to place the scales in a parallel position and, move the stage micrometer until the starting lines of both scales coincide (Figure 2).  Find another point, as far along the scale as possible, where two other division lines are exactly superimposed.

Figure 2.      Alignment of the ocular micrometer and stage micrometer.

Left: ocular micrometer; right: stage micrometer.

5.  Move the 40X objective into place (so 400X magnifications).  In order to determine the length that is equivalent to one division on the ocular micrometer scale, count the number of divisions on the ocular micrometer and the corresponding length on the stage micrometer scale (each division of the stage micrometer measures 0.01 mm  ). You can now calculate the width of one division on the ocular micrometer since you know how many ocular divisions are equivalent to one stage division.

6.  Remove the stage micrometer.   Obtain two  different slides of cells, H9c2 cells (a permanent  cell  line  derived  from  embryonic  BD1X  rat  heart  tissue)  stained  with Hematoxylin-eosin stain and human blood smear (contains erythrocytes, leukocytes and platelets) stained with Wright's stain.

7.  Place the slide of cells on the microscope stage and focus on the cells to be measured. Superimpose the ocular micrometer scale and count the number of ocular division equivalent to the cell diameter (choose the largest dimension).  Multiply the number of divisions by the width of single division to give the actual diameter.

8.  Measure the length or diameter of five different cells for H9c2 cells and one specific type of leukocyte.   Calculate the average diameter.   Record all data on  Datasheet accordingly.

Note: Use the SAME magnification (400X) in calibrating the ocular micrometer to measure the cell diameter.  You may use 100X for ocular calibration but then you have to use the same magnification to measure cell diameter.

E. Drawing morphology of H9c2 cells and leukocytes

With the given slides, observe and draw the respective morphology of H9c2 cells and leukocytes.

1.  Place the slide of cells on the microscope stage and focus on the cells under 40X high-dry* objective,

*High-dry means high power magnification by non-oil immersion lens, which is the 40X objective in the microscope of BIO2032.

2.  Draw the morphology on the Datasheet. Label nucleus, nuclear membrane, cytoplasm, and cell membrane in your drawings.

F. The hemocytometer

In many cases it is important to know how many cells one has in a suspension. Using a special device called a hemocytometer.  Each hemocytometer has two separate counting areas.  Each counting area has 9 large squares (see Figure 3).  In the figure:

A is a large square with 16 small squares;

B is a large square with 20 small squares, each with 4 rectangles;

C is a large square with 25 small squares, each with 16 smaller squares.

A special coverslip is supported above the grid such that the volume above each large square is equal to precisely 10-4 ml (equivalent to 0.1 μl).  If 30 cells were located in one large square, then your sample would contain 30 x 104 cells/ml.

The trick in using a hemocytometer is in getting an even distribution of cells over the squares. To do this,first the hemocytometer with its coverslip in position is placed on the microscope stage.  A drop of evenly suspended cells is then introduced to the edge and fluid is drawn in by capillary action.  Note the sample CANNOT be used under the following circumstances:

-     the volume is too great and the fluid flows into the gutters,

-     it takes more than one application to cover the grid,

-     the drop is held too long before applying it to the hemocytometer.

To check on accuracy, the numbers from several squares are obtained.  If they are close, they can be averaged.  If the numbers are far apart, the hemocytometer should be cleaned and reloaded.  A comparison of the numbers from several loadings should be made.

Figure 3.       Diagram of the grids on a hemocytometer cell counting chamber G. Determining cell concentration by using the hemocytometer

1.    HL-60   cells   (human   leukemia   cells)   in   cultured  medium  are  provided   in  a microcentrifuge tube with unknown cell density.

2.    Obtain uniformly suspended cells by inverting the tube several times.

3.    Load both counting chambers with the cell suspension using a micropipette and tip. Approximately 10 μl will be required for each chamber.   Place a coverslip over the counting chambers of the hemocytometer allow the cell suspension to fill the space (some hemocytometers may contain a V-shaped groove for loading).

Note: Fill the entire volume of the chamber but do not overfill!

4.    Wait for a while until the cells stop drifting around inside the chamber.

5.    Determine the  number  of cells by viewing the cells under a microscope at  100x magnification.  Count all of the cells contained within the four large squares labeled as A in Figure 3 (count only those cells on the lines of two sides of the small squares to avoid counting cells twice). You are required to count both of the chambers on the hemocytometer.

Note: Always lower the stage before inserting a hemocytometer.   It is much thicker than any ordinary microscope slide.

6.    Calculate the cell concentration (number of cells/ml) on the Datasheet accordingly.

Experimental Datasheet of Exercise 2

I.  Measuring the field size

Diameter of the 100X field (by calculation) =                                               mm or                                        μm

II.  Ocular calibration and cell diameter measurement

l  Calibration of the ocular micrometer : 1 ocular division =                        mm

l  Average length of H9c2 cell  =                                 μm

l  Average diameter of                  cell  =                                μm

III.  Examining two cell types

Drawing morphology

IV.  Determining cell concentration of HL-60 cells

l  Average cell number of HL-60 suspension  =                               X 104 cells/ml

Please show all of your calculation steps.