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Posted in General

Haemocytometers: You Can Count On Us!

Apart from the incubators and the Class II hoods, perhaps one of the most overlooked but important pieces of equipment in the cell culture lab is the haemocytometer. For accurate cell-seeding experiments and splitting cells, this thick glass slide is an essential tool for everyone working with live cells.

There are a number of different haemocytometers on the market and each one has a different grid pattern as well as different recommended uses.

Louis-Charles Malassez (1842-1909) was a French histologist and anatomist and is the person credited for inventing the haemocytometer. As the name suggests, this device was originally intended for the quantitative counting of blood cells.

The most frequently used haemocytometer is the Neubauer (or ‘Improved Neubauer’) chamber. Other haemocytometers include the Burker, Thoma and Fuchs-Rosenthal. Most of the haemocytometers are manufactured from crystal glass and generally measure 30 x 70 mm with a thickness of 4 mm. Two vertical lines are ground from the glass to define the counting area and the double cell counting chambers have a ground out ‘H’ shape. A selection of haemocytometers is available from Agar Scientific;

http://www.agarscientific.com/lm/slides-coverslips/haemacytometers.html

 

The Neubauer/Improved Neubauer haemocytometer

The cell counting area of the Neubauer measures 3 mm2 (with each of the main squares measuring 1 mm);

A1

The grid is subdivided and the 16 squares at each of the four corners measure 500 mm2. In a Neubauer chamber, the central square consists of 16 smaller squares each one with the same width/height the corner squares. Each of the 16 central squares is further subdivided into 16 smaller squares. The glass covers for haemocytometers are specifically designed with regards to thickness and size (generally 22 mm2). Do not be tempted to use a coverslip as this may result in an inaccurate cell count. When the coverglass is placed over the counting area, this leaves a specific area for which to introduce the cells/liquid to be counted. The Neubauer chamber is designed to leave a gap of 100 mm between the top surface of the counting area and the bottom surface of the coverglass.

The Improved Neubauer has a slightly different grid pattern compared to the ‘old’ Neubauer chamber. The overall size of the central counting area is still the same, but the square in the central cross area are divided into 25 smaller squares, each one measuring 400 mm2;

A2

Each of the 25 squares in the centre of the cross grid are further subdivided into 16 smaller squares which measure 2.5 mm2 (or 0.0025 mm2 as etched onto the surface of the Improved Neubauer slide). In other words, the central cross square contains 400 small squares.

The Neubauer chambers are designed with these grid patterns for the counting of blood cells (although, these chambers are suited for the counting of most mammalian cell lines/primary cells etc.). The larger squares at the four corners are designed for the counting of white blood cells whereas the central smaller squares are for counting red blood cells and platelets.

 

The Burker Haemocytometer

The Burker chamber has the same dimension of grid (and same depth) as the Neubauer slide;

A3

The main difference is that the central square is not sub-divided into the 16 or 25 smaller squares. The grid pattern runs across the whole counting area and the central square contains the same number of smaller squares (16) as the corner and side squares.

The Thoma Haemocytometer

The Thoma chamber has the same dimensions and depth as the Neubauer chamber, however, this haemocytometer does not have the four larger corner squares;

A4

The Thoma haemocytometer seems to be the favoured counting chamber for microbiologists.

The Fuchs-Rosenthal Haemocytometer

This counting chamber is specifically designed for the counting of cells from cerebro-spinal fluid. The main difference (apart from the counting grid pattern) is that the distance between the coverslip and the chamber surface is 200 mm. This gives a larger volume of cells/liquid which can be counted and is also used for counting blood cells.

A5

The grid pattern of the Fuchs Rosenthal is similar to the Burker chamber, except that the Fuchs Rosenthal haemocytometer is designed to accommodate twice as much volume compared to the other chambers mentioned above.

Counting cells using a haemocytometer

For this example, we will consider the counting of a sample of cells using an Improved Neubauer haemocytometer (although this can be applied to any of the haemocytometers above excluding the Thoma chamber).

Firstly, you’ll need to clean and prepare the chamber- the easiest way is with 70% ethanol. Dry the chamber (preferably with a lens cleaning tissue). You will then need to moisten the edges of the chamber and the most common way is with the exhaled breath with mouth fully open. Slide on the coverglass using gentle pressure and you should see something called ‘Newton’s Rings’ where the coverglass is in contact with the chamber (rings caused by an interference pattern).

  • Prepare the cell suspension

The cells and fluid (such as culture medium, PBS etc.) should be well mixed to provide a homogenous sample. You can do this by gently pipetting the cells up and down a pipette tip or by gently agitating the flask/container. Don’t be tempted to use excessive pipetting or a vortex mixer as this may shear cells.

Quickly remove a volume of cells and transfer to an Eppendorf tube. Add an equal volume of trypan blue and gently mix by pipetting or by tapping the tube (note- this is an example using a 1:1 dilution, but you may need to adjust depending on the cell density). Using trypan blue will help to distinguish between dead (blue) cells and living (clear) cells.

  • Loading and counting cells
  1. With a pipette, carefully draw up around 20 ml of the cell mixture. Place the pipette tip against the edge of the coverglass and slowly expel the liquid until the counting chamber is full. Capillary action will help to ensure that the counting chamber is full, but care should be taken not to overfill the chamber. A volume of 10 ml is sufficient to fill one counting chamber.
  2. Place the haemocytometer on the microscope stage. Using the 10X objective, focus on the grid lines. The general rule for counting of cells which fall on the edges of the squares is only to count those cells which are on the top and the left hand side grid lines. This rule ensures that you do not count the same cells twice.
  3. It is advisable to use a hand tally to keep count of cells. Count the number of cells in one of the corner squares (which are further separated into 16 squares). Count all of the clear cells within the squares and those touching the top and left hand grid lines. Another tip is to count in a ‘turret’ pattern, i.e., count along the top row of squares from left to right, down to the next square and count from right to left and so on until all the cells in the 16 small squares have been counted.
  4. Move to the next corner square and repeat until you have counted the four larger corner squares.
  5. The volume of liquid between the coverglass and the counting chamber is such that the number of cells which are counted in one set of 16 small squares is equivalent to the number of cells X 104/ml.

 

  • Calculating the total number of cells

Add the four counts together and divide by four to give an average over the whole counting chamber.

Multiply by 2 to take into account the 1:1 dilution made when adding the trypan blue to the cell mixture.

The total count from 4 sets of 16 corner squares = (cells/mL x 104) x 4 squares from one hemocytometer grid.

In other words;

Total cells (X104)/ml = Total cells counted x (dilution factor/number of squares counted)

For example;

-Total cell count was 180

-Number of squares counted was 4

-100 ml of trypan blue was added to 100 ml of cell suspension

Therefore;

Total cells (X104)/ml = 180 x (2/4)

Which gives 90 X 104 cells/ml.

If you find that there are too many cells in your suspension, then simply increase the dilution factor of cells to trypan blue (but remember to take this into account in the final calculation of cell density).

For an accurate determination of the total number of cells, the number of cells in one of the large squares should be between 15 and 50.

 

AUTHOR: Martin Wilson

Posted in General

Finder Slides: Finding and Re-Locating Areas of Interest on Microscope Slides

How often have you been scanning a new batch of slides and found an area of interest which you wish to show to a colleague or come back to at a later date? Marking the slide or the coverslip with a marker or an engraver is impractical as this could easily crack the coverslip, or obscure the area of interest under a blob of ink.

This is where the ‘Finder Slides’ are invaluable accessories. These slides have a grid pattern of co-ordinates usually over the entire surface of the slide. They are not overlaid on top of the specimen slide, instead, the area of interest is found on a slide which is then carefully removed without disturbing the XY stage controls and replaced with a finder slide. Looking back down through the eyepieces, you will then see co-ordinates which relate to the area of interest.

 

Maltwood’s Finder (1858)

One of the earliest examples of a finder slide was the ‘Maltwood’s Finder’. This finder slide was invented by Thomas Maltwood (1827-1921), a fellow of the Royal Microscopical Society. These early finder slides were manufactured by Smith, Beck & Beck of London. James Smith started making microscopes in 1839 and went into partnership with Richard Beck on 1847. Four years later, Richards’s brother Joseph joined the firm. Smith, Beck & Beck started making the Maltwood’s Finder slides in 1858.

Maltwood initially drew the grid on a sheet of paper measuring 10 inches square which was photographed to give a negative measuring one inch square. A positive microphotographic plate was printed from this original negative and hand-mounted on a slide measuring 3” x 1 ¼”. On this plate the lines were only 1/50th of an inch apart with a total of 2,500 co-ordinate squares. These glass slides were hand engraved with the manufacturer names and each one was numbered.

 

Webb’s Finder (1880)

William Webb (1815-1888) is well known amongst antique microscopy enthusiasts for his microscopic writings etched onto glass slides. Webb had constructed a micro-engraving machine which reduced his hand movements to incredibly small scales, but the details of the machine are lost and it is thought that he destroyed it shortly before his death. Amongst the many curious slides which he made were epigrams measuring only 1/5000th of an inch and the entire Lord’s Prayer etched into glass and measuring 1/500th of a square inch. He proposed that these slides were “the best, the most simple, and unerring tests for objectives”. In 1880 he put his micro-engraving machine to more practical use and he produced finder slides and he boasted that these were of greater accuracy that the Maltwood’s Finder slides in that the Webb’s Finder contained 16 squares in the space of one square of a Maltwood’s Finder.

 

Gage Finder/Marker (1895)

Simon Henry Gage (1851-1944) was an American microscopist and Professor of anatomy, embryology and histology at Cornell University. In 1895, he published a paper in the Proceedings of the American Microscopical Society and he describes the problems of finding exact points on a specimen slide; “in one’s private study or for exhibition to friends, the special point that is on a slide is often the last to be found, much to one’s discomfiture.” This device differed from the classic finder slides as it actually marked the coverslip surface with a delicate brush filled with “colored shellac or other varnish”. This apparatus replaced one of the objectives and when an area of interest was found, the apparatus was swung into position and the marker turned to make a circle around the area of interest.

 

The England Finder

Today, one of the most common finder slides is ‘The England Finder’ which is available from Agar Scientific;

http://www.agarscientific.com/the-england-finder.html

The England Finder is not named after the country, but after its inventor, Charles Norman England. He devised the original layout of the finder over 50 years ago and working with a company which was called ‘Graticules Ltd.’, they created a photomask layout as a freehand plot. Graticules Ltd. later became ‘Psyer-SGI Ltd.’ but the mastering of each of these slides is still derived from the original freehand plot which not only makes each Finder slide identical, but also a very unique product which cannot be reproduced by modern methods.

In a letter dated 1959 to the New Scientist from the Quekett Microscopical Club, Graticules Ltd. were described (in relation to the Maltwood’s Finder) as having “performed a public service in reintroducing an accessory of considerable usefulness in microscopy.”

The England Finder has a vacuum-deposited chromium grid which covers the entire slide surface and is particularly suited for lower-power magnification work. Finder slides mean that the microscopist can record the co-ordinates of any area of interest on a microscope slide which can then easily be found at a later date or by a colleague using the same (or a similar microscope). The England Finder can be used on any microscope with an X-Y stage movement of 75 mm by 25 mm. The grids line are situated at 1 mm intervals- along the X axis, the grids are labelled 1 to 75, whilst the Y axis grid lines are labelled A to Z (excluding ‘I’ to avoid confusion with the number ‘1’);

This gives a total of 1875 positions across the whole slide. In addition, each co-ordinate is divided into five (central circle and four quartiles) for even greater positioning accuracy (there are 9045 locations on each slide).

When using a finder slide, it is also useful to have a cross-hair graticule placed in one of the eyepieces of the microscope for even greater accuracy. Selections of such graticules are available from Agar Scientific;

http://www.agarscientific.com/eyepiece-graticules-e-f.html

To use the England Finder;

  • Firstly mark the slide orientation (X/Y) on the label or frosted section of the specimen slide. Make sure that the microscope stage is referenced to its maximum X and Y stop positions.
  • Locate the centre of the area of interest under the cross-hairs of the graticule.
  • Being careful not to move the position of the microscope stage, carefully remove the specimen slide and replace with the England Finder making sure the X/Y orientation is the same as the specimen slide.
  • Bring the England Finder grid into focus and make a note of the numbers and letters which are under the cross-hairs. Record the letter and numbers of the main circle followed by the number of the quartile (if the exact spot falls into one of these sections). For example, ‘U39/1’. If the point of interest lies in the centre of one of the circles, then this should be recorded as ‘0’.
  • To find an area of interest from a co-ordinate, ensure the stage is correctly referenced to X and Y, place the England Finder on the stage and locate the co-ordinate, then carefully replace the finder slide with the specimen slide. The area of interest should now be in the centre of the field of view.

AUTHOR: Martin Wilson

Posted in General

Here Comes The Sun: The Science of the Summer Solstice

sun

Given the weather in the UK during most of May, you could be forgiven for thinking it was still spring (although, technically it was)! However, Sunday 21st June 2015 sees the summer solstice dawn upon us. So, what exactly is the summer solstice and have we really approached midsummer already?

The word ‘solstice’ derives from the Latin ‘sol’ meaning ‘sun’ and ‘sistere’ meaning ‘to stand still’. This standing still refers to the path of the sun and ‘declination’ which is a means of measuring the angle of a celestial body on the ‘celestial sphere’. This is an imaginary sphere around earth upon which all objects in the sky can be thought of as projected onto the underside of the sphere or dome. Similar to the latitude and longitude system used here on earth, it is a means by which astronomers can describe the position of stars and planets relative to a co-ordinate system. Declination specifies a position in the sky which is relative to the equator and the poles.

The earth doesn’t spin on an axis which is vertical, instead, the angle of tilt of the north/south axis is approximately 23.40 and this gives rises to the seasonal variations we experience on earth relative to the amount of sunlight we receive. At the Northern Hemisphere during the summer solstice, the North Pole is tilted towards the sun.

 

Factors affecting the exact date

The way in which a year is measured is not exactly an accurate science- it’s why we have leap-years. A year actually last for 365.24 days, so every four years we catch up with the extra day on February 29th. This means of measuring time also has an effect on the dates and exact times of each of the solstice and equinox dates. This year, it’s the 21st June, whereas in 2016 it will fall on the 20th June. The last time there was a summer solstice on the 22nd June was back in 1971.Other factors also influence the exact timing of the summer solstice. For example, the earth does not rotate at a constant speed around its elliptical orbit.

 

Axial precession

Another factor which occurs over a vast time scale is known as the ‘precession of the equinoxes’ (or ‘axial precession’). The earth is spheroid in shape and not a prefect sphere- indeed the equatorial diameter of the earth is 43 Km larger than the polar diameter. As described above, the earth it tilted in its orbit and this means that the ‘bulge’ around the equator is off-centre relative to the gravitational pull of the sun. This results in a small amount of torque as the gravitational force of the suns pulls harder on one side of the earth relative to the distant side. It’s not just the sun which exerts a gravitational pull on the earth- our own moon and other planets in the solar system also exert a pull on us. If the earth was a perfect sphere, precession would not occur.

If you extend an imaginary line through the centre of the earth (titled at 23.40), then this would trace a circle as the earth moves through the cycle of precession. One complete precession cycle takes 25,772 years. This cycle changes the exact times and dates of the solstices and equinoxes. Another interesting point is that the exact positions of stars in our sky also change through the cycle of precession. Polaris is our current ‘Pole Star’, however, 13,000 years from now, this will change and Vega (in the constellation of Lyra) will become our Pole Star. Polaris will get another turn at being the Pole Star in approximately 26,000 years from now!

 

Midsummer? But we’ve only just started!

The exact date of the solstice and what we know as midsummer do not fall upon the same days. Although midsummer festivities and celebrations are pagan in origin and would have fallen on the equinox days, the Christian church took over this festival and designated the date as the 24th June and is associated with the nativity of John the Baptist.

However, midsummer doesn’t mean the middle of summer. In meteorological terms, summer starts with the equinox and lasts for the months of June, July and August. In astronomical terms, the solstice marks the beginning of summer and it ends with the autumnal equinox on September 23rd. The average highest temperatures in the UK are usually in the months of July and August. This is partly due to the way that earth retains heat from the sun and the fact that we are an island. The oceans surrounding us act as a huge ‘heat sink’, absorbing and re-radiating the heat from the sun. Although the earth absorbs the most intense of the sun’s rays around the date of the solstice, it takes several weeks for the heat to be released, hence the fact that our hottest days are usually in July or August. If there were no oceans on earth, our hottest days would fall around the date of the solstice.

Depending on the location on earth, the summer solstice is generally regarded by those of us living in the Northern Hemisphere as the ‘longest day’. At the South Pole there will be 24 hours of darkness, whereas the North Pole will receive 24 hours of daylight. At the equator, there is approximately 12 hours of daylight. Here in the UK, the day length varies with London seeing around 16 hours and 38 minutes of daylight at the solstice, whereas in Edinburgh, the day length is approximately 17 hours and 36 minutes. Up in Shetland, they will see a day as long as 18 hours and 55 minutes.

 

Megalithic monuments and the solstice

Perhaps the most famous of the megalithic sites in the UK which is associated with the solstice is Stonehenge, although there are many other ancient sites in this country and around the world which are aligned to astronomical events.

Archaeologists have found evidence of very early wooden structures at the Stonehenge site which date back around 10,000 years ago. Even then, the post holes were aligned east/west which may have signified a link to the movements of the sun.

The earliest monumental building phase at Stonehenge was around 3,100 BC and consisted of a 110 metre circular bank and ditch with a gap facing the north east. The latter stages of monument building at the site maintained this NE/SW alignment. It was the English antiquarian and archaeologist William Stukeley (1687-1765) who first noted and recorded the rising of the summer solstice sun above the Heel Stone in the summer of 1720. The sun doesn’t rise exactly above the Heel Stone when viewed from the centre of the circle, but this is unsurprising when taking into account the vast time period and factors such as precession. Evidence has also shown that the Heel Stone was only one of a pair of such stones and the solstice sunrise would have been framed by these stones at the time of the building.

After much controversy and legal battles, English Heritage now grant access to Stonehenge at the times of the solstices and equinoxes (the stones are roped off to visitors at other times of the year);

http://www.english-heritage.org.uk/visit/places/stonehenge/plan-your-visit/summer-solstice/#

Author: Martin Wilson

Posted in General

Love in the Lab: Why Sexism is Still an Issue

women in science

Sexism in science reared its ugly and unwelcome head again recently when Sir Tim Hunt stood up on the 9th of June to address the World Conference of Science Journalists in Seoul, South Korea. In front of an audience of science journalists from around the world, he reportedly said the following comments;

“Let me tell you about my trouble with girls. Three things happen when they are in the lab: you fall in love with them, they fall in love with you, and when you criticise them they cry.”

He later gave an interview for the ‘Today’ programme on BBC Radio 4 stating that he did “mean the part about having trouble with girls”, but went on to try to apologise for his comments adding that he was “really sorry that I said what I said”, and that it was “a very stupid thing to do in the presence of all those journalists”. He had hoped his remarks were “intended as a light-hearted, ironic comment” but instead they were “interpreted deadly seriously by my audience”. Is the joke still funny when no one else is laughing?

Let me tell you about my trouble with these comments. Firstly, I have never worked in a laboratory which has employed ‘girls’ (or boys for that matter). Although laboratory visits from schoolchildren should be encouraged, they shouldn’t be made to don lab coats and crack on with an RT-PCR, well, not at least until they have left school! Many women view the word ‘girl’ as derogatory and its use in this context can imply emotional and intellectual immaturity.

Secondly, as humans, we all feel a huge range of emotions, from love to hate and everything in between. However, to let our emotions towards fellow scientists somehow hinder or cloud the science we are employed to do seem to hint at a lack of professionalism. Because of our emotions and judgements, there will undoubtedly be colleagues we like and those we dislike. But respect is the key and regardless of our feelings towards other scientists, if they are good at their job, then surely that is the fundamental issue.

Despite Tim Hunt’s comments, should we have rushed to judge him and publically head-hunt him through social media and the press? I’m not defending his personal viewpoints, but how would it have been perceived if a senior female scientist had made such comments about ‘boys’ in the lab? Would there have been such a media outcry? Unfortunately, there is still a huge gender gap between senior scientists; only 16 % of full-time professors in the STEM subjects (science, technology, engineering and maths) are women.

Tim Hunt is 72 and he attended a single-sex school in the 1960’s. His own views about ‘girls’ in the lab may be part of the conditioning from that era and from his educational environment. In his statement he also said “my trouble with girls”, implying this was a personal issue and problem for him. Indeed, in an exclusive interview with Tim Hunt and his wife, Professor Mary Collins in The Observer newspaper 1, Collins stated that “really it was just part of his upbringing.” Although she also admitted that “it was an unbelievably stupid thing to say”.

Shortly after the conference, a rather vicious flurry of social media comments ensued and he was described on Twitter as a “clueless, sexist jerk”. Yes, his personal viewpoints are sexist, but a clueless jerk? Tim Hunt was a Fellow of the Royal Society, but they immediately distanced themselves from his comments and in their statement said that “Too many talented individuals do not fulfil their scientific potential because of issues such as gender”. On the 11th June, the Royal Society issued a statement to say that Tim Hunt had resigned 2. They stated that although he had “made exceptional contributions to science in terms of his own research on the cell cycle and its implications for our understanding of cancer”, his “recent comments relating to women in science have no place in science.” The final comment in their statement is the most telling; “It is the great respect that he has earned for his work that has made his recent comments so disappointing”.

The Royal Society works to promote diversity in science and has many projects, awards and campaigns targeted not only at gender inequality in science, but also ethnicity, disability and those who may need flexibility in their work due to roles as carers and parents 3. Tim Hunt should have been aware of the work of the Royal Society in his professional role before making seemingly personal comments which contradict the vision of diversity promoted by the Royal Society.

Furthermore, Tim Hunt was an Honorary Professor with University College London (UCL) Faculty of Life Sciences- that is until 10th June 4. In the interview in The Observer, his wife was informed by UCL that he had to resign or face the sack (Tim Hunt was still flying back from South Korea at this point). With regards to his resignation, the university stated that “UCL was the first university in England to admit women students on equal terms to men, and the university believes that this outcome is compatible with our commitment to gender equality.”

What have these thirty seven words from Tim Hunt done to highlight gender inequality in science? They have shown that even at the highest level of academia, intelligence and (supposed) professionalism that such views still exist. It may partly be a generational issue, but inequality exists in our laboratories, institutes and universities. In real terms, this manifests as the fact that there are still far fewer women working in the STEM subject areas and these women are paid less than their male counterparts. More worryingly, one study examining trainees in scientific fieldwork has reported that women are 3.5-times more likely to experience sexual harassment than men 5. Whilst Tim Hunt’s statement was disappointing, it highlights  the very real issues of gender inequality in science and rather than head-hunting an individual, we should focus our attention on addressing the root causes with a view to achieving gender neutrality in science.

1 http://www.theguardian.com/science/2015/jun/13/tim-hunt-hung-out-to-dry-interview-mary-collins

2 https://royalsociety.org/news/2015/06/sir-tim-hunt-resigns-from-royal-society-awards-committee/

3 https://royalsociety.org/about-us/diversity/

4 http://www.ucl.ac.uk/news/news-articles/0615/100615-tim-hunt

5 http://journals.plos.org/plosone/article?id=10.1371/journal.pone.0102172

Posted in General

Helping Cells and Sections to Stick: Cleaning, Sterilising and Coating Slides and Coverslips

Although we very briefly touched on slide coating in a previous article, I wanted to describe in more detail the many ways in which slides and coverslips are prepared for different purposes including growing cells on them and helping sections to stick to them through the process of immunohistochemistry (IHC).

In this article, we will look at cleaning and rinsing of slides/coverslips, sterilising slides/coverslips for growing cells and different methods of coating slides/coverslips. There is no single correct way to prepare slides or coverslips and it will depend on factors such as the cell-line being used.

 

Cleaning slides and coverslips

Although many slides are supplied as pre-washed (such as these ones http://www.agarscientific.com/lm/slides-coverslips/microscope-slides.html ), many labs and scientists still prefer to wash their own slides/coverslips and it is obviously more important to do so if using items from a packet which is already open. A freshly opened box may look clean, but there may be a thin film of grease from the manufacturing process which will prevent optimal adherence of cells.

  • > Washing with PBS/water

Although many protocols advocate washing in water or PBS before use, such washing may not get rid of any oil/grease based films from the surface of the glass and the addition of PBS may in fact leave salt crystals on the surface when it dries.

  • > Acid Washing

Acid washing of coverslips is recommended particularly if you are planning to grow adherent cells on the surface and it helps polypeptides to adhere to the glass. It is a relatively long protocol though, so it is best to plan in advance and make up a batch of acid-washed coverslips and store them in a clean container.

  1. Separate the coverslips and incubate them in 1M HCl at 500C to 600C for between four and 16 hours.
  2. Allow the HCl to cool to room temperature and then rinse two times in double-distilled or ultrapure water.
  3. Add double-distilled or ultrapure water to the coverslips and sonicate for 30 minutes. Repeat this twice using fresh water each time.
  4. Sonicate the coverslips for 30 minutes in a solution of 50% ethanol (make up all the ethanol solutions with double-distilled or ultrapure water).
  5. Sonicate the coverslips for 30 minutes in a solution of 70% ethanol.
  6. Sonicate the coverslips for 30 minutes in a solution of 95% ethanol.
  7. Remove the final ethanol wash and store the coverslips in 95% ethanol until use.

A slightly shortened version of this protocol involves taking the coverslips from step (3) above, rinsing them in 100% ethanol, drying and storing them in a clean airtight container until use. Other similar methods involve the use of commercial glass cleaners in place of the hydrochloric acid.

 

Sterilising coverslips

  • > Ethanol

Most protocols for sterilising coverslips advocate the use of ethanol in concentrations ranging from 70% to 100%, either with or without further sterilisation steps.

If using ethanol alone, the coverslips can be placed in the bottom of a sterile 6-, 12- or 24- well plate and covered with 70% ethanol, left for five minutes and repeated three times. Follow this with a final wash in 100% ethanol.

A more thorough method (and more time consuming) is to dip a coverslip in 70% ethanol and flame with the blue flame of a Bunsen burner. There’s an obvious hazard here though- keep the ethanol stock well away from the flame! Flaming will eliminate most micro-organisms and spores from the surface of the glass.

  • > UV Light

UV light is often used in conjunction with 70% ethanol. Dip the coverslips in 70% ethanol and then expose to the UV light in a tissue culture hood for between 20 and 30 minutes. Alternatively, UV exposure can be used on its own.

  • > Autoclaving

This is one of the easiest methods to sterilise coverslips. Simply place the coverslips in a glass petri dish and send through the dry cycle of an autoclave. This is typically a 20 minute step. Some protocols advocate a 70% ethanol wash beforehand, but autoclaving alone will ensure that the glass is tissue culture sterile. Just remember to open the petri dish in a tissue culture hood and use sterile forceps to remove each one.

 

Coating slides and coverslips

As with cleaning and sterilising, there are many methods for the coating (or ‘subbing’) of slides and coverslips. The coatings used depend largely on the cells which you wish to grow on the glass. The most common coating is poly-lysine, but we will also take a look at collagen, laminin and fibronection.

  • > Poly-Lysine

Poly-Lysine enhances the electrostatic interaction between the positively charged ions on the surface of the glass and the negatively charged attachment ions of the cell membrane. Poly-Lysine not only increases the electrostatic bond between cells and the glass surface, but also helps to ensure that frozen and paraffin embedded sections remain stuck to slides during the many IHC steps.

Two types of poly-lysine are available and the choice will depend on the cells type you are using. Poly-L-lysine is the most commonly used and is also suitable for tissue sections. However, some cell lines can excrete proteases which break down Poly-L-lysine, so it is recommended to use Poly-D-lysine instead.

Poly-L-lysine has been shown to be a suitable coating for cells such as primary neurons, neuronal cell lines, PC12 cells and HEK293 cells. As with many of these cleaning and coating methods, there are a variety of options, but below is a standard method for poly-lysine coating;

  1. Clean the slides/coverslips before coating. If you are coating coverslips to grow cells on, then it is recommended to acid-wash these beforehand.
  2. Make a 1 mg/ml poly-lysine solution in sterile water. If coating coverslips, place these in a sterile plastic petri dish and cover with poly-lysine solution. If coating slides, place in a clean slide rack in a dish and again, cover with poly-lysine solution. The optimum ratio of poly-lysine solution to glass surface is 1 ml: 25 cm2 (this works out around 900 slides/litre of solution).
  3. Place slides or coverslips on a rocker and rock gently for 5 minutes at room temperature.
  4. For coverslips, remove solution by aspiration and wash with sterile water. Allow to dry for at least 2 hours before use. For slides, pour off poly-lysine and dry in a 600C slide oven for an hour.
  5. Coated glass can be stored at 40C in a sterile container for up to three months.

Pre-coated poly-lysine slides are also commercially available, such as these ones from Agar Scientific;

http://www.agarscientific.com/lm/slides-coverslips/polysine-microscope-slides.html

  • > Collagen

Collagen is an extracellular matrix protein which can be used on coverslips to promote adherence of cells such as epithelial and endothelial cells as well as cells lines such as CHO and HEK293 cells. The most commonly used collagen is type I (isolated from rat tails). Other types used for adherence include type II and type IV, but also use the recommended type depending on the cells you wish to grow on the coverslips.

  • > Fibronectin

Fibronectin is also an extracellular matrix protein which contains an RGD sequence (Arg-Gly-Asp) which mediates cell attachment. For this matrix protein, the species source is unimportant as all contain the RGD sequence. Many cell types will attach to fibronectin coated glass including endothelial cells, fibroblasts, neuronal cells, and CHO and HEK293 cell lines.

  • > Laminin

Laminin is an extracellular matrix protein and is a major component of one of the layers of the basement membrane. Laminins bind to cell membranes through integrin receptors and other plasma membrane molecules. Laminins are a suitable coating and substrate for a wide variety of cell types ranging from fibroblasts to neuronal cells. Some protocols advocate pre-coating of slides/coverslips with poly-lysine before laminin coating for extra adherence.

 

AUTHOR: Martin Wilson

Posted in General

The Origins and Development of the Confocal Scanning Microscope

It all starts in New York City in the summer of 1927 with the birth of Marvin Lee Minsky. Following his high school education (and a brief stint in the US Navy during WW II), he went on to gain a BA in mathematics from Harvard, and a PhD in the subject from Princeton before joining MIT in 1958. He founded the MIT Computer Science and Artificial Intelligence Laboratory is Professor of Electrical Engineering and Computer Science. In Isaac Asimov’s autobiography, he admitted that there were only two people he considered to be more intelligent than he was, one was Carl Sagan, and the other was Minsky. In addition, Minsky was also an advisor on the movie ‘2001:A Space Odyssey’.

 

The need to see neurons

In Minsky’s memoir on inventing the confocal scanning microscope, he explained that his home was always full of prisms, optics and lenses- his father was an ophthalmologist. At a young age, he started taking optical systems apart and rebuilding them. At both Harvard and Princeton, he studied biology, neurophysiology and neuroanatomy in addition to his mathematical studies. At the time, not much was known about how brains function and how nerve cells were connected. Minsky admits to being frustrated by this lack of understanding and although the shapes of many nerve cells were visible, what was needed was a three dimensional ‘wiring diagram’. However, with the imaging systems at the time (and given the sheer density of the nerve cells of the brain), it was impossible to distinguish such neural networks due to the scattering of light. Minsky knew that what was needed was an instrument capable of optical sectioning to eliminate the out of focus light.

 

Zirconium arcs and a military surplus radar

Whilst studying at Harvard, he was fortunate enough to be given a room in the physics laboratory with permission to use and buy whatever equipment he needed. Minsky designed his symmetrical microscope with an objective lens and a pinhole at either side of the specimen to eliminate the scattered and out-of-focus light. When he made his invention, there were no lasers suitable to illuminate the specimen with the intensity of light needed, so he used zirconium arcs instead which was a time-consuming process- each point scan taking up to 10 seconds. The final image was produced on a military surplus radar scope which he had acquired.

The next design problem he faced was whether to move the specimen or move the optics for the scanning procedure. Minsky admitted that the prospect of moving two tiny pinholes simultaneously daunted him, so he decided to keep the optics in a fixed position and move the stage instead. He machined all of the parts for the prototype himself, spending months in the machine shop at Harvard- the skills he learnt during this time also helped him to design and build a robotic hand and arm 10 years later.

 

1955: The first patent for a scanning microscope

Were it not for the fact that his brother-in-law was a patent attorney, Minsky’s invention may never have been documented at all. Minsky admitted that he didn’t keep notes or write down what he was working on! On November 18th 1955, he sent a patent letter entitled ‘Double focussing stage scanning microscope’ and the confocal was born.

 

1966-1967: A Nipkow disc confocal

In 1966, a patent was filed by two Czechoslovakian scientists, Mojmir Petran and Milan Hadravsky for a tandem-scanning microscope. This confocal used a Nipkow Disc. Briefly, this is a mechanical spinning disc which has a spiral pattern of thousands of individual pinholes drilled in it. The effect of the Nipkow disc is that thousands of points on the specimen are illuminated simultaneously. This was actually the first commercially available confocal scanning microscope being sold by a small company in Czechoslovakia and by Tracor-Northern in the US. The first scientific paper describing the instrument and its uses was published in the journal Science in 1967. However, the instrument only worked well for the brightest of specimens.

 

1969: The first laser scanning confocal

The first true laser scanning confocal microscope was designed and built by M. David Egger and Paul Davidovits from Yale University who published a paper on their findings in Nature in 1969. This used a 633 nm He-Ne laser and opposed to the earlier confocal systems (and the next one in the time-line) the sample remained stationary and was illuminated by the movement of an objective lens.

 

1977-1979: Naming the confocal

It wasn’t until 1977 that the term ‘confocal’ was used to describe such microscopes in a publication on theoretical analysis in ‘Optica Acta: International Journal of Optics’ by two scientists from Oxford, Colin J. R. Sheppard and A. Choudhury. The term ‘confocal’ means ‘having the same or common focus’. There is another claimant to using the term ‘confocal microscope’ for the first time- a Dutch physicist called G. Fred Brakenhoff developed a laser scanning microscope in 1979. He published his findings in the Journal of Microscopy and it seems to be the first time the term Confocal Scanning Light Microscope (or ‘CSLM’) was used.

Most of these early confocals proved to be too inconvenient for biological applications being both too slow and too sensitive to vibrations.

 

The 1980’s: Time to buy!

In 1982, a company called ‘Oxford Optoelectronics’ (since acquired by Bio-Rad) offered the first commercially available stage scanning CSLM which was connected to a computer (the ‘SOM-25’). This was commercialised from a design by a team based at Oxford University.

In 1986, at the Medical Research Council Laboratory of Molecular Biology in Cambridge, a team of scientists were busy working inside a tent made from World-War II blackout material. Inside the tent, a new prototype CSLM capable of scan speeds in excess of 4000 lines per second was being developed. This instrument used a galvanometer-driven mirror for frame-scanning and a rotating polygonal mirror for line scanning. The line scanning mirror was proving to be problematic and producing chromatic aberrations when green and red fluorophores were used. The polygon was subsequently replaced with two oscillating galvanometer mirrors. Further developments were made to allow adjustable scanning as well as dealing with the chromatic aberration and the patent for the design has since been used by Bio-Rad for all of their confocal point-scanning systems. The team had approached other leading microscope manufacturers including Zeiss and Leica, but this proved to be unfruitful. However, a commercial agreement was soon concluded with Bio-Rad and the ‘MRC 500’ was launched to an excited audience in 1987.

It wasn’t long until the other main microscope manufacturers were developing and building commercially available instruments, either based upon the MRC 500, or in parallel to this microscope. The development of the confocal microscope continues and in the years since the MRC 500 was launched we have seen major advancements including multi-photon techniques and the ability to image 90 separate colours in a single sample. It is an exciting and rapidly expanding field- watch this space!

AUTHOR: Martin Wilson

Posted in General

Microscope Configurations: A Brief History of the Compound, Inverted and Stereo Microscopes

It is still unclear when the first microscope was invented and who was credited with the invention. The general opinion is that a Dutch spectacle/eyeglass maker by the name of Zaccharias Janssen (1585-1632) invented and constructed a microscope sometime between 1590 and 1595. However, this ‘general opinion’ doesn’t add up- if Janssen was born in 1585 and he invented and constructed the first microscope, then he would have done so at the age of five! Another theory alludes to the later date of 1595 and that his father helped to construct the instrument. Or that he was born in 1580.

This fist microscope was a simple sliding tube giving between approximately 3X and 10X magnification. The eyepiece consisted of a bi-convex lens (i.e. both surfaces of the lens are convex) whereas the objective lens was a plano-convex lens (i.e. one flat and one convex side). Another claimant to the title of ‘inventor of the microscope’ was a German born spectacle maker called Hans Lippershey (1570-1619) who actually lived next door to Janssen in Middelburg in the Netherlands.

Then there’s Galileo! In 1609, Galileo Galilei (1564-1642) built a device which he named the ‘occhiolino’ (little eye). Giovanni Faber (1574-1629) was an anatomist, botanist and close friend of Galileo. They were both members of an Italian science academy (‘Accademia dei Lincei’) founded in 1603. Faber was very excited by Galileo’s new instrument and he is the person responsible for naming this instrument as the ‘mircosope’  from the Greek words ‘micron’ (meaning ‘small’) and ‘skopein’ (meaning ‘to look at’).

 

The history and mystery of the Leeuwenhoek microscopes

Antonie van Leeuwenhoek (1632-1732) is commonly known as the ‘Father of Microbiology’, but he also made many advances in the field of microscopy. Whilst working as a draper, he developed an interest in lens making. He was making tiny high-quality glass spheres which he used as the lenses in his microscopes. It is thought that he made around 25 microscopes and it is claimed that these offered magnification in the range of between 275 and 500X. Leeuwenhoek kept his microscope construction and lens making techniques very secret.

Mystery still surrounds his microscopes and there was an excellent article by Brian J. Ford in Laboratory News from February 20151 regarding some of his instruments which have recently come up for auction and subsequently disappeared. One was bought at Christies auctioneers in 2009 for over £300,000 supposedly by a representative of a European biosciences organisation. This was allegedly going to go on public display, but the microscope has disappeared. Then in 2014, a collection of items appeared on eBay which were dredged from a canal at Delft. The seller described one of the items as a ‘weird drawing instrument’, however, it was soon clear that this appeared to be an original Leeuwenhoek microscope. Shortly after, the items were removed from eBay and the vendor disappeared. Apparently a Spanish collector had offered to buy the whole lot for 1,500 Euros, but he never received the items and his money was refunded. It remains to be seen if these hugely important instruments ever turn up again.

 

A need for binocular vision

Throughout the hundreds of years of development of the instrument, microscopists had always desired to view specimens through a binocular set-up. The first stages of development of this configuration were not entirely satisfactory as the splitting of the final image resulted in halving the numerical aperture of the objectives. Ernst Leitz II (1871–1956) was the second son of the entrepreneur Ernst Leitz who founded the ‘Ernst Leitz Optical Works’ at the beginning of the 20th Century (later to become ‘Leica’). Ernst Leitz II realised that a physical beam-splitter was needed in a binocular configuration which would ensure that the performance of the objectives was unaffected when the image was split. This binocular tube design soon became the standard for all such configurations and the principles are still used to this day.

 

Inverted microscopes

The history of the inverted microscope is slightly more recent. John Lawrence Smith (1818-1883) was an American chemist and during his time as Professor of Chemistry at the University of Louisiana (which is now Tulane University), he invented the inverted microscope in the year 1850. He published this work in the American Journal of Science and Arts in 1852 with a paper entitled ‘The Inverted Microscope- a new form of Microscope; with the Description of a New Eye-piece Micrometer and a New Form of Goniometer for Measuring the Angles of Crystals under the Microscope’2.

Although the modern inverted microscope is primarily used for viewing live cells and tissue (usually in flasks or dishes), the invention came about to facilitate the micro-chemical research which Smith was undertaking at the time. He describes the problem in his paper;

“The great obstacle to chemical research beneath the microscope are two-fold; first, the necessity of manipulating in the limited space between the object-glass and the stage; and, secondly, the exposure of the most essential parts of the instrument to the vapors emanating from the re-agents employed, and the condensation of vapor on the under surface of the object-glass souring the view. A less important obstacle is the impossibility of heating a liquid or other substance while beneath the microscope. The only way by which these difficulties can be surmounted is to place the object-glass beneath the stage, and the object above it, with an optical arrangement of such a nature as to permit observation.”

In others words, the heating of chemicals on the stage would produce vapours which would lead to corrosion of the objectives! Although he alluded to the fact that his invention may have other uses, he was more concerned with heating and evaporation of chemicals than the viewing of living systems.

 

Stereo microscopes

Stereo microscopes are also known as stereoscopic or dissecting microscopes which is a more descriptive name for their purpose. The first pseudo-stereoscopic microscope was designed and built by a monk called Cherubin d’Orleans (1613-1697). In 1677 he made a small microscope with two separate eyepieces and objective lenses, though at the time, he was unaware of how the perception of depth was created by his invention. In wasn’t until 1852 that the English scientist and inventor Charles Wheatstone (1802-1875) first described the principles of stereoscopic vision. He published this as a paper in the Philosophical Transactions entitled ‘On Some Remarkable, and Hitherto Unobserved, Phenomena of Binocular Vision’3.

Around the same time, a further advancement in stereo microscopy was discovered by an American scientist called John Leonard Riddell (1807-1865). He published his findings in the Quarterly Journal of Microscopical Science in 1854 in his paper ‘On the Binocular Microscope’4. The difference with Riddell’s configuration is that the microscope used a prism system with a single objective- usually referred to as Common Main Objective (CMO) stereo microscopes. However, with the Riddell instrument, the image formed was reversed (and therefore still ‘psuedo-scopic’).

In the early 1890’s, an American biologist and instrument maker, Horatio S. Greenough developed a stereo microscope which was an alternative design to the CMO microscope. Much like the design which Father d’Orleans had made, Greenough’s microscope used two separate but identical optical paths. He approached the Carl Zeiss Company with his design where the Zeiss engineers changed the plans slightly- Greenough had designed a lens erecting system to ensure correct orientation of the final image, but at Zeiss, this was replaced with image-correcting prisms. To this day, stereo microscopes are still based on the ‘Greenough principle’.

 

  1. http://www.labnews.co.uk/features/deepening-mystery-of-disappearing-microscope/
  2. https://archive.org/stream/mobot31753002152418/mobot31753002152418_djvu.txt
  3. http://rstl.royalsocietypublishing.org/content/142/1.full.pdf+html
  4. http://jcs.biologists.org/content/s1-2/5/18.full.pdf+html

AUTHOR: Martin Wilson

Posted in General

Science and Politics: An Election 2015 Special

Science

Please find below a summary of our Science and Politics: An Election 2015 Special  document.

You can download the full document at the bottom of this blog for free.

In summary:

  • >>The Conservatives promise to train an extra 17,500 maths and physics teachers over the next five years, but aren’t they forgetting about biology and chemistry?
  • >>The Green Party are ‘pro-science’, but are calling for an immediate end to all experiments using genetically modified (GM) organisms (GMO)- where will this leave the current and future disease models?
  • >>The Labour Party would introduce a new long-term funding policy framework for science and innovation, however, being their manifesto doesn’t mention ‘agriculture’ or ‘farming’- what would become of GM foodstuffs research under a Labour government?
  • >>‘Britain will be the place to be if you want to thrive in science’, say the Liberal Democrats. But is their environmental science policy to only allow Ultra-Low Emission vehicles to use UK roads by 2040 over-ambitious?
  • >>Plaid Cymru are calling to make Wales, the UK and the EU a GMO-free zone- will this affect our food, or will it extend to biological research?
  • >>Innovation Centres in Scotland have been set up by the SNP to foster research, innovation and commercialisation, but their manifesto only mentions ‘science’ on one occasion. How do they plan to keep the pipeline of future scientists flowing?
  • >>UKIP want to rejuvenate and expand the UK coal industry and coal-fired power stations. What will this mean for air pollution and the Climate Change Act?

DOWNLOAD THE FULL DOCUMENT HERE FOR FREE ( No details required.)

Agar Scientific. Science and Politics. An Election 2015 Special

 

Posted in General

Microscopes – They Don’t Clean Themselves!

There are three things which most research grade and laboratory microscopes share in common: (1) They are relatively expensive pieces of equipment, (2) They are delicate instruments with a complex series of components and (3) They don’t clean themselves!

One of the most important aspects of the use, care and maintenance of a microscope is keeping it clean. There’s the financial investment issue in the instrument as well as the time investment in preparing specimens and slides. The last thing you want is for dust and dirt to contaminate your images. Prevention is better than cleaning and the easiest way to keep a microscope clean is to use the dust cover when the instrument is not in use. If you don’t have a dust cover, you can always buy one, or even use a large thick bin liner or bag- anything which prevents dust from settling on the lenses.

It should be emphasised that improper cleaning of lenses and parts can cause as much (if not more) damage than leaving a neglected instrument. For example, tiny grit particles can cause scratching on the surface of lenses, cleaning solvents can affect coating on the lens surface and if too much solvent is used, this can lead to degradation of the cement which is used to hold objective front lenses in place.

If you can see dust/dirt/smudges or blurring when looking down the eyepieces, the first thing to do is locate the source of contamination. This can be done by checking each of the objectives in turn. If the contamination remains, then it is likely to be on the eyepieces/sub-stage condenser or field diaphragm. For routine cleaning, these are the only components which should be considered. Do not attempt to clean any of the internal optical components of a microscope unless you know exactly what you are doing! This is best left to a specialist as part of the maintenance contract, or to a knowledgeable microscope technician.

There are four items which you should invest in before tackling the cleaning of a microscope- an air duster, lens cleaning tissue, cotton buds (or ‘Q-tips’) and a lens cleaning solvent/solution.

 

Air Dusters
Two types of air dusters are available for cleaning optics- aerosols and bulb/bellows dusters. If using an aerosol, it is important to use one which is specifically manufactured for cleaning optics, i.e. one without an oil-based propellant. Do not shake the can before use and always use whilst holding upright- this prevents the freezing liquid propellant from being sprayed out along with the air. Don’t be tempted to blow off dust particles as you will inevitably blow a fine mist of saliva over the optics to complicate the issue.

For an air duster which is safe to use on microscope optics, be sure to have a look at ‘Dust-Off Plus’ which is available from Agar Scientific;

http://www.agarscientific.com/dust-off-plus.html

 

Lens cleaning tissue/cotton buds
Although general lab wipes and tissues can be used for cleaning the microscope body and stage, these should never be used to clean lenses. Lens cleaning tissues are designed to be less abrasive than lab wipes as well as being tightly woven to prevent fibres being left on the optics after use.

Two types of lens cleaning tissue are available from Agar Scientific- optical lens tissue and the well-known Whatman 105 lens cleaning tissue;

http://www.agarscientific.com/optical-lens-tissue.html

http://www.agarscientific.com/lens-cleaning-tissue.html

If using cotton buds in addition to tissue, it should be remembered that many of the commercial cotton buds will not be 100% cotton and may contain synthetic fibres which can cause damage to lenses. However, there is much contradictory advice online regarding cotton buds. Some sites recommend synthetic materials as they regard pure cotton to be abrasive, whilst other sites extol the virtues of pure cotton as they regard synthetic buds to be too harsh for delicate optical surfaces! Personally, I would recommend high quality cotton wool wrapped around bamboo splints or cotton swabs- both of which can be obtained from Agar Scientific;

http://www.agarscientific.com/cotton-swab-pointed-double-head-7-6-cm-box-200.html

http://www.agarscientific.com/cleaning-materials.html

 

Cleaning solvents
Before using any cleaning solvents or solutions on optical components of the microscope, you should always check the manufacturer’s recommendations. The solvents described below are for information only and it is important to determine suitability before use.

Xylene is widely used for cleaning lenses, but care should be taken with its use due to potential toxicity. It can also leave a small residue where the last of the solvent evaporates off the lens. Similarly, both ethanol and isopropanol/isopropyl are also popular cleaning solvents.

Many microscope facilities have their own cleaning solvent ‘recipe’. However, if you are starting work with your own microscope, then a cheap alternative is a commercial window cleaning solution.

Most of the major microscope companies have their own propriety brands of optical cleaning solutions.

Finally, distilled water should be the first solution applied to a dirty lens (unless you are certain that the contamination is oil/grease based). Solvents should be the last resort!

 

Removing immersion oil
In one of my previous articles, I shared some practical tips for the use and cleaning of immersion oils;

http://www.agarscientific.net/looking-down-and-through-microscope-optics-3-oil-immersion-objectives/

To briefly recap, immersion oil should be removed from the objective as soon as possible after use. If residual oil is left on the lenses, a small amount of xylene can be used to remove it. Don’t use water, acetone or alcohol to try to remove immersion oil as it’s not soluble in any of these.

 

Steps for cleaning

  1. Locate the source of contamination.
  2. If you need to remove the optical component, then wear a pair of powder-free gloves (this also protects you if you are using a solvent-based cleaning fluid). You don’t want to cover the other parts of the microscope with fingerprints whilst you clean a dirty component!
  3. Place a clean piece of lab roll/tissue on the bench on which to place the component(s). If you need to remove the eyepieces, be sure to cover the eyetube with a tissue to prevent dust from settling in the tube.
  4. Use an air duster to remove non-attached dust. If the source of contamination is water soluble, use this first. Do not soak or spray microscope components in water or any cleaning fluids. Always wet the cotton bud/lens tissue in a small amount of fluid. Never use a dry cotton bud or lens tissue on any optical lens.
  5. If you need to remove an objective from the nosecone, then place it screw side down on the clean tissue. With a damp cotton bud, slowly start in the centre of the lens and in a circular movement, work your way out towards the lens rim. Working in a zig-zag movement will only serve to spread the contamination. Do not use excessive pressure to try to remove dirt/smudges- pressure applied to a front lens can knock it out of alignment, or produce tiny gaps in the cement though which fluids/immersion oil can penetrate. Do not try to clean the back lens of an objective. Although you can air-dust this part if necessary, cleaning inside an objective is something which should only be done by a specialist or knowledgeable microscope technician.
  6. Of all the components of a microscope, the eyepieces are likely to be the source of contamination. Carefully remove the rubber eyecup surrounding the eye lens before cleaning the lens itself. Clean the eyecups separately using a small amount of lab detergent in water, but make sure they are completely dry before placing back on the eyepiece.
  7. If the sub-stage condenser or the field diaphragm is the source of contamination, then make sure that the diaphragm is fully open before cleaning. These delicate parts are easily damaged and fibres from the cotton bud/tissue could become trapped in the diaphragm.
  8. To clean the actual body or stage of the microscope, you can use a lint-free cloth or tissue and a small amount of lab detergent and water, or you can also use a small drop of the window cleaning spray.
  9. For each round of cleaning, always use a new cotton bud or tissue. It is false economy to re-use these for the cleaning of expensive and delicate optical components.

With all of these suggestions, always follow manufacturer guidelines before cleaning any component.

AUTHOR: Martin Wilson

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Posted in General Tagged , , , , , , , ,