Tag Archives: cells

Meeting with Ashish Mehta – iPSCs


I had a preliminary meeting with Dr Ash Mehta on Monday (21/2) to discuss the iPSC generation using blood. He uses the ThermoFisher CytoTune-iPS Sendai Reprogramming system and has generously offered to guide me through the cell reprogramming/iPSC generation timeline.  The great thing about doing this is that the cells are technical immortal when in a stem cell state.  This will enable me to achieve the primary project aims even if immortalisation of primary cells via SV40 does not work.

We followed up again on Tuesday (22/2) to go over the process in more detail and set up basic project requirements including blood collection in collaboration with the clinical research team. As part of this process, Ash introduced me to a lovely phlebotomist who agreed to collect my blood, as well as the Menzies Clinical Research Facility Manager to ensure everyone is informed about the project and correct processes are in place to move forward. After supplying project documentation and confirmation of ethics clearance, I received final sign off from the Chair o CRFMC to proceed with blood collection on 23/2/ – so full steam ahead!

Ash also showed me around his lab and allowed me to view the PMBCs (peripheral blood mononuclear cells) he thawed last week. The cells are cultured in suspension (non-adherent) and are circular in shape.


PBMCs in culture – image courtesy of Ashish Mehta 

He also showed me some iPSCs and the difference between stem cell colonies and cells that have started to differentiate.

iPSC Colonies

iPSC colony. Image courtesy of Ashish Mehta

About PMBCs

As part the introduction to cell reprogramming, Ash explained the basics and value of working with blood cells.

Blood is made from a number of different cell types including red blood cells (erythrocytes), white blood cells (macrophages, lymphocytes, monocytes, neutrophils, eosinophils, basophils ) and platelets (thrombocytes). Platelets and red blood cells have no nuclei so they cannot be reprogrammed and only the mononucleated (single nucleus) white blood cells are suitable for the process.

To isolate PMBCs, the blood undergoes  gradient centrifugation which separates the blood into layers of cell types via density.

Gradient Separation

Diagram of peripheral blood separated into different layers including PBMCs ( round cells with a single nucleus: lymphocytes, monocytes, natural killer cells (NK cells), dendritic cells).

The advantage of using PBMCs is that you can tell more readily when the virus has initially successfully reprogrammed cells, as they change from non-adherent to adherent and start forming dense colonies of small cells.

The colonies need to be maintained meticulously as they tend to differentiate in culture (i.e. turn into (uncontrolled) specific cell types).

Ash indicated that when the blood is collected, it should be processed (PBMCs extracted) within a 4-hour window. A vial of blood should yield 4 – 5 million PBMCs, so he suggested that we freeze 4 x vials (1 x 106) as backup and proceed with a single 1 x 106 sample. This will also need careful planning to ensure that I am able to donate and process blood on the same day, plus move forward with the next steps involved.

On Thursday 24/2, Ash has kindly agreed for me to shadow him when he adds the Cytotune 2.0 (Sendai Virus reprogramming system) to the cultured PMBC samples. I’m looking forward to learning more.


Media Change and Cell Maintenance

While I wait for ethics clearance and team members to return to the lab, I am continuing to culture my cells.

Cell Culture 21/1/22

3 x T75 and 1 x Cut Glass Dish ready for cell maintenance (media change) on 21/1/22.

At this point, I am feeding more confluent cells on a weekly basis (depending on how they look):

T75 Flask #1- 21/1/22T75 originally plated 15/12/21 – Flask #1

T75 Flask #1 - 21/1/22T75 originally plated 15/12/21 – Flask #2

T75 Flask #3- 21/1/22T75 originally plated 7/10/21

Cut Glass Dish - 21/1/22Cut Glass Dish – originally plated 15/12/21

While the cut glass dish is hard to image due to the more uneven surface of the glass, and being contained in a larger Petri Dish, there seems to be a good level of cell growth. Perhaps this will result in better staining in the next round.

I still have four other T75s. However, since these are less dense in cell numbers, I am limiting media change to reduce ongoing maintenance costs and media usage:

T75 P4T75 originally plated 7/10/21 and passaged 16/11/21 – Flask #1

T75 P4 - Flask #2T75 originally plated 7/10/21 and passaged 16/11/21 – Flask #2

T75 P5T75 originally plated 15/12/21 – Flask #3

I must admit that I am continually amazed that the original flask of cells plated by Jo-Maree in August last year (and passaged a month later) still has viable cells in it. While much of the T75 flask area is pretty sparse with cells, there are some larger clusters including a growing bunch of elongated fibroid-like cells:

Org plated cells 1T75 originally plated 10/09/21 showing sparse number of cells with evidence of previous cell movement and presence in ‘ghost trails’.

Org plated cells 2T75 originally plated 10/09/21 showing small cluster of cells with evidence of previous cell movement and presence in ‘ghost trails’.

Org plated cells 3T75 originally plated 10/09/21 showing larger cluster of fibroid-like cells.

They are easy to miss, but I’ve made a note to monitor their progress and see how they proliferate. Again…it just takes one mutant to get a cell line going!


H & E staining is a bust :(


After spending most of the day in the lab staining up the cut glass dishes and vessels…

Staining set up

… I have emerged victorious-less.

Cut Glass after Staining

Cut Glass after Staining Microscope images of cut glass dishes after H&E staining on 20/01/22 showing scratches on glass surface and no cells. 

There are no cells visible at all – just scratches on the surface of the glass. This is likely due to the very limited number of remaining cells which may have been further dislodged during the washing process.

The glass vials have not fared much better. While there are cells visible, most of them are dead or dried out as it was tricky working with the small opening and 3D surface area.

Cell Vial 1Microscope image of glass vial after H&E staining on 20/01/22. The image shows a vast number of dead cells that were not fixed in a live state. 

Cell Vial 2Microscope image of glass vial after H&E staining on 20/01/22. The image shows dried cell remnants. 

The flasks show relatively good fixation of the cells, but the staining is not really visible under the light microscope in the ‘dirty’ lab.

T25 - Fixed

Microscope image of fixed PHGL Tumour Baby Cells after H&E staining on 20/01/22. The image shows intact  fixed cells with very limited evidence of H&E stain.

I think, I will stick to Petri Dishes for the next test as they offer a more consistent surface area to work with.

Review of Cell Flasks

Despite overall loss of cells, one of my P4 flasks which had a good level of cell growth previously still has a strong number of cells.  It also shows ‘ghost trails’ over a prolonged period of culture without passage. The flask was originally plated out at P3 on 7/10/21 passaged on 9/11/21 with remaining cells maintained every week.

P3 Cell Flask Image of PHGL Tumour Baby cells Flask with original plating date (P 3, 07/10/21) and passage date (P4, 09/11/21) recorded. 

Images taken 15/12/21:

PHGL TB P3 at 15/12/21Light microscope image of PHGL Tumour Baby P3 in T75 flask [Org plated 07/10/21] imaged on 15/12/21.

PHGL TB P3 at 15/12/21

Images taken 13/1/22:

Tumour Baby P3 - 13/01/22

Tumour Baby P3 - 13/01/22

Tumour Baby P3 - 13/01/22Light microscope images of PHGL Tumour Baby P4 in T75 flask [Org P3 plated 07/10/21 – passaged P4 on 9/11/21] imaged on 13/1/22 after cell maintenance. 

I maintained the cells (media change) on 13/1/22 and will allow them to grow for another week before I passage them and set up new experiments with cut glass and Petri dishes.

Prior to leaving for the festive season, I passaged my more confluent culture plates to establish fresh T75 flasks.

Flask #1

Tumour Baby P5 Flask 15/12/21Photograph of PHGL Tumour Baby P5 in T75 flask plated on 15/12/21 prior to holidays. 

Images taken 16/12/21:

Tumour Baby P5 16/12/21

Tumour Baby P5 16/12/21Light microscope image of PHGL Tumour Baby P5 in T75 flask plated on 15/12/21.

Images taken 13/1/22:

Tumour Baby P5 13/1/22

Tumour Baby P5 13/1/22Light microscope images of PHGL Tumour Baby P5 in T75 flask plated on 15/12/21 and imaged on 13/1/22. 

Flask #2

Tumour Baby P5 Flask 2 15/12/21Photograph of PHGL Tumour Baby P5 in T75 flask #2 plated on 15/12/21.

Images taken 16/12/21:

Tumour Baby Flask 2 P5 16/12/21Light microscope image of PHGL Tumour Baby P5 in T75 flask 2 plated on 15/12/21.

Tumour Baby Flask 2 P5 16/12/21Light microscope image of PHGL Tumour Baby P5 in T75 flask 2 plated on 15/12/21.

Images taken 13/1/22:

Tumour Baby Flask 2 P5 13/1/22

Tumour Baby Flask 2 P5 13/1/22Light microscope image of PHGL Tumour Baby P5 in T75 flask 2 imaged on 13/1/22.

In both flasks here is some cell growth present. However, much less than expected. This may be due to my very low seeding ratio which was deliberate to avoid over-confluence during my absence. I changed the media for both flasks on 13/1/22 and will continue to maintain them to see if there is any further growth.

Ghost Movement Trails in Original Tumour Baby Cell Flask

As indicated in my first post-holiday post, my flasks mostly still have live cells – although some of the more confluent flasks also resulted in mass cell death. Interestingly, my first thawed flask of cells – with remaining cells maintained after the first passage – still has living cells after four months of continuous culture:

P1 Flask - 9/10/21Light microscope image of PHGL TB cells at 13/1/22 showing a main cluster area of remaining cells. 

These cells were originally thawed and plated on 10/09/21. After passage on 21/09/21 the original flask continued to be maintained with a weekly feeding/media change regimen.

P1 TB FlaskPhotograph of P1 PHGL TB flask originally plated out by Jo-Maree on 10/0921. 

Tumour Baby P1 21/09/21Light microscope image of P1 PHGL TB cells taken on 21/09/21 prior to passage. 

Tumour Baby P2 23/09/21Light microscope image of P2 PHGL TB cells (in original flask) taken on 23/09/21 after cell passage on 21/09/21. A few cells remain visible in the flask. 

At present, here are only very small clusters of cells remaining:

Tumour Baby original flask viewed 13/1/22Light microscope image of P2 PHGL TB cells (in original flask) taken on 13/1/22. The image reveals a ‘sprinkle’ of cells beyond the main cluster. 

The rest of the flask is filled with the ghostly trails of cellular movement:

Trails of Cell Movement

Trails of Cell MovementLight microscope images of P2 PHGL TB cells (in original flask) taken on 13/1/22. The image reveals trails of cellular movement and existence. 

Since this is pretty consistent with prolonged culture, I am curious to show Jo-Maree to determine what the ‘residue’ is. It is quite poetic to consider the way in which  traces remain of different movements and interactions. I have decided to continue to maintain the flask until there are no more viable cells, so it will be interesting to see how these traces evolve.


Extent of death post-holiday – Cut Glass and Petri Dishes

Following more detailed review of cell flasks, there are some hardy ‘survivors’ of my lab-free holiday.

Cut Glass CellLight microscope image of cut glass dish with media containing dead cell debris and evidence of a small number of surviving cells. 

After removing the old media, it was easier to see the remaining cells:

Cell Survivors - Cut Glass

Light microscope image of cut glass dish in PBS showing evidence of a small number of surviving cells. 

This image shows even more evidence of cell survival:

Cell Survivors - Cut GlassLight microscope image of cut glass dish in PBS showing further evidence of a small number of surviving cells. 

 Cell Survivors - Cut GlassLight microscope image of cut glass dish in PBS with likely cells circled. There are a few additional potential cells visible, but I have only circled the most obvious. 

To get an even better sense of survivors, I will fix (in 4% PFA) and H&E stain 2/3 of the cut glass dishes. The flatter cut glass dish and Petri dish (with more potential for cell survival will be maintained in the incubator to see how they fare over the next week).

Petri Dish Light microscope image of Petri dish with fresh complete media with live cells. 

I will also fix and stain the glass vessels. It is less likely that these will yield anything interesting, but it will help me troubleshoot how to do the protocol with the tiny openings – it is very difficult to effectively remove the media – even with a 20ul pipette and tip 🙁



Immunostained Cells

The lab has a great set-up for fluorescence microscopy which makes imaging quick and easy.

You just need to load the well plate into the machine and set up basic imaging parameters. You do need to image both DAPI and Phalloidin stains, but the software merges the images for you.

Fluorescent Images of Fibroid Cells

Task Manager

Loading ModeSimple graphic interface with presets ready to complete fluorescent microscopy. 

As discussed in my previous post on immunostaining, the blue dots indicate nuclei and the green structures reveal the cytoskeleton via binding to actin.

Confluent wells: 

DAPI & Phalloidin PHGL Tumour Baby Cells

DAPI & Phalloidin PHGL Tumour Baby Cells

DAPI & Phalloidin PHGL Tumour Baby Cells

DAPI & Phalloidin PHGL Tumour Baby Cells DAPI and Fluorescein Phalloidin staining of confluent fibroid cells P4 (although this is potentially misleading as the cells are very slow growing). 

Less confluent wells:

DAPI & Phalloidin PHGL Tumour Baby Cells

DAPI & Phalloidin PHGL Tumour Baby Cells

DAPI & Phalloidin PHGL Tumour Baby Cells

DAPI & Phalloidin PHGL Tumour Baby Cells

DAPI & Phalloidin PHGL Tumour Baby Cells

DAPI & Phalloidin PHGL Tumour Baby Cells DAPI and Fluorescein Phalloidin staining of fibroid cells P4  which enables better visualisation of individual cells. 

Immunostaining Protocol

It is time to complete the immunostaining protocol with guidance from Jo-Maree. I must admit that with holidays looming, my note taking was a bit sketchy. I will need to follow up with Jo-Maree to record the correct details of DAPI and Fluorescein Phalloidin stain. This will ensure that I know how to prepare (and order) stocks in the future.

I already prepared a couple of wells at different cell concentrations ready for staining.  We had to delay the protocol, so some of the higher concentration wells are likely a bit over-confluent. It will be interesting to see how they look under the fluorescence microscope.

CELL FIXATION: ‘Dirty’ Biolab

Working in Fume HoodWorking with 4% PFA in Fume Hood

Prior to imaging, I fixed the cells in 4% PFA:

  1. Remove culture media (discard in waste container with bleach)
  2. Wash cells with PBS (discard in waste container with bleach) x 2
  3. Move cells to fume hood
  4. Add 4% PFA to each well for 15 – 20min at room temp (in fume hood)
  5. Remove 4% PFA solution (discard in PFA waste container in fume hood)
  6. Add PBS (make sure cells are covered or they dry out and produce poor images

IMMUNOSTAINING: At lab bench area

Lab BenchWorking at lab bench in the Stroke Group area

  1. Remove PBS from each well
  2. Add 1mL 0.3% Triton X-100 (a strong detergent) to permeabilize cells (make cells permeable – this allows the phalloidin stain to enter the cell structure) for 10 min
  3. Make up DAPI (5mL PBS Tween and 1ul DAPI) and protect from light with aluminium cover
  4. Remove Triton X-100 and add 1mL DAPI solution to each well and incubate at room temp (with aluminum cover to protect from light) for 5 minutes
    Aluminium Cover
  5. Make up Flouroscein Phalloidin (1mL PBS sand 2μL Flouroscein) and protect from light
  6. Remove and discard DAPI solution
  7. Add 1mL PBS 0.1% Tween to each well for 5 min then discard x 3 (i.e. wash with PBS Tween x 3)
    PBS Tween
  8. Add Phalloidin stain to each well.
  9. Incubate at room temp for 1 – 2 hours
  10. Remove Phalloidin solution
  11. Wash with PBS Tween x 2
  12. Add 1mL PBS in each well
  13. Cover with aluminum foil to protect from light.
  14. Cells are ready for imaging. They can be stored in the fridge (with aluminium foil cover) until ready.


Immunostaining of fibroid cells

To gain some more insight into the cellular structure of my fibroid cells, I asked Jo-Maree to help me with immunostaining. Immunostaining refers to a staining method that uses antibodies to stain different proteins and structures within the cell. We are going to start with an easy protocol using two antibodies: DAPI and Phalloidin.

DAPI (4′,6-diamidino-2-phenylindole) is a very common fluorescent blue stain used to reveal the nucleus in cultured cells. It can penetrate the intact membrane of the cell. Thus, it can be used for staining both fixed and live cells.

Microscopic image of stem cells, Hues 9 stained with DAPI (blue)Microscopic image of stem cells, Hues 9 stained with DAPI (blue) by the UC San Diego Stem Cell Program.


  1. Add DAPI to the complete culture medium used for cultured cells at a concentration of 10 ug/mL.
  2. Remove culture medium from the cells and replace with the medium containing the DAPI.
  3. Cover cells from light exposure and incubate at room temperature or 37°C for 5-15 minutes, then image.

Direct Addition: According to the biotium protocol, you can also stain cells by adding the dye directly to the cell culture and medium. However, this requires a 10X concentration of dye. 

  1. Add the dye to complete culture medium at 10 times the final recommended staining DAPI concentration – 100 ug/mL..
  2. Without removing the medium from the cells, add 1/10 volume of 10X dye directly to the well.
  3. Immediate mix thoroughly by gently pipetting the medium up and down. For larger well sizes (e.g., 24-well to 6-well plates), the plate can be gently swirled to mix.
  4. Cover cells from light exposure and incubate cells at room temperature or 37°C for 5-15 minutes, then image


  1. Add DAPI to PBS at 1 ug/mL.
  2. Add the PBS with dye to cells or tissue sections and incubate at room temperature for at least 5 minutes with covering from light exposure.
  3. Samples can be stored in a lightsafe covering (e.g. aluminium foil) at 4°C after staining and before imaging

From: https://biotium.com/tech-tips/protocol-staining-cells-with-hoechst-or-dapi-nuclear-stains/ with minor edits for simpliticy.

Since we are using phalloidin and fixing the cells prior to imaging, we need to follow some additional step to make the cells permeable.

Phalloidin Jo-Maree did not have any relevant stocks, Natalie kindly sourced a sample from another group. [I am always impressed by the generosity of functioning labs and how groups are happy to share stocks to enable other researchers to move forward.]

We are using Fluorescein Phalloidin as a counterstain to enable the visualisation of actin – a protein found in large quantities in the cytoskeleton and cell muscle fibres. As such, it  plays a vital role in cell muscle contraction and overall cell movement.

Interestingly Phalloidin is a toxin (specifically phallotoxin) derived from Amanita phalloides (death cap mushroom).

Amanita phalloidesImage of Amanita phalloides via Wikimedia Commons

Phalloidin binds very well to actin filaments and is therefore very useful in visualising cell structure.

Phalloidin staining of actin filamentsU2OS cells stained with fluorescent phalloidin taken on a confocal microscope by Howard Vindin


  1. Fix cells in 3–4% formaldehyde in PBS at room temperature for 10–30 minutes.
  2. Remove fixation solution and wash cells 2–3 times in PBS.
  3. Add 0.1% Triton X-100 in PBS into the fixed cells for 3–5 minutes to increase permeability. Then wash cells 2–3 times in PBS.
  4. Add phalloidin-conjugate working solution. Incubate at room temperature for 20–90 minutes.
  5. Add DAPI DNA staining dye at this point.
  6. Rinse cells 2–3 times with PBS, 5 min per wash.
  7. Cover with lightfast material to preserve fluorescence

Adapted from:  https://www.abcam.com/protocols/phalloidin-staining-protocol 

Hematoxylin and Eosin Staining

Jo-Maree finally had some time to go over basic H&E staining procedures. Since my HBVPs are fixed on the base of  glass Petri Dishes, the process is much less involved than working with wax embedded specimens.

H&E is a very common stain combination used in histology. Hematoxylin stains nuclei blue-purple
Eosin stains cytoplasm (protein, muscle fibres etc.) pink
H & E Stain Protocol Basic H&E staining protocol from Jo-Maree.   We only need to follow the staining process.

Stain: washing Petri Dish on bench in Histology Lab at MSP with Erlenmeyer flask containing distilled water for washing. 

Prior to adding the Hematoxylin stain, we washed the Petri dishes with distilled water (DW). Usually, we would simply wash the dishes under running water from the tap. However, since rapid water could dislodge the cells from the base of the dish, we have used a beaker to control the water flow.  I washed each dish twice to remove PBS and dislodged cells.

Hematoxylin StainHematoxylin Stain – deep red stain 

Contrary to what the name Hematoxylin suggests, the dye is actually naturally derived and comes from the tree  Haematoxylum campechianum (Logwood). As such, it is non-toxic and does not need to be added in a fume cabinet. The dye was added to the Petri Dishes for 5 mins, then washed with distilled water.

The next step involved adding ammoniated water (approx 2 – 3 drops ammonia to 400mL distilled water) to the stained cells for 30 secs.   This process is referred to as ‘bluing’ and helps change the red – purple hematoxylin to a blue – purple color.

Hematoxylin Stained DishCells visible on the base of Petri Dish following Hematoxylin staining.

After washing the Petri Dish thoroughly after ‘bluing’, we added the Eosin stain.  Eosin is a xanthene dye and has an intense fluorescent colour.

Eosin StainEosin stain in Petri Dish.

The Eosin stain only needs 2 mins to stain the cytoplasm and matrix of cells. Following  another thorough wash of the dish, we added 95% ethanol and secured the Petri dish lids with parafilm.

For stained sections on glass slides, it is usual to add Xylene (toxic) and a coverslip. In this case, we could either create large scale glass covers (a bit impractical) or clear resin. I think clear resin is the best solution as it would create a barrier and preserve the dyed cells. I am keen to use the fixed cells in dishes as part of sculptural works.  However, I will need to check with lab manager David Steele that I am able to remove these fixed cells from the lab.