Protocol for imaging micro beads?

[stu_the_flat]

Hi

I'm new to this forum. I have tried searching this forum but I can't find a topic. I apologize if I missed it.

I'm a PHD student I'm relatively new to confocal imaging I am trying to image sub resolution beads to measure the point spread function.

I am using a bio rad system I use a 60X water immersion lens I set the system to 1024 X 1024 at 16 bit resolution. Kalman sampling was set to 4 and the scanning head was as slow as possible and the laser power was as low as possible to measure the beads.

I have found that this is the perfect recipe for measuring photo bleaching!

I tried looking around for some sort of protocol on imaging sub resolution beads but I was unable to find anything. Also as I understand it there will always be some element of photo bleaching? Is it easy to compensate for if I can calculate how much beaching as occurred.

Similarly if anybody as any tips on creating the slides with the beads I would be grateful mine are quite messy.

Many thanks

Stuart McIntyre

[M. van de corput]

Before using the beads stock solution sonicate the tube shortly to
destroy bead aggregates.

__
This is how I make my bead slides. Some do not use a coating but my
beads always started swimming without coating._
_Mariette Kemner-van de Corput
Erasmus MC, Rotterdam, NL

_
_

_TetraSpeck beads: http://probes.invitrogen.com/media/pis/mp07279.pdf _

200nm TetraSpeck 4 colour polystyrene beads (Invitrogen)

500nm TetraSpeck 4 colour polystyrene beads (Invitrogen)

ex/em 365/430; 505/515; 560/580; 660/680

 

100nm green fluorescent polystyrene beads (Duke Scientific) for blue,
green and red beads

 

Use the same chemicals, microscope slides and cover slips as you use in
the experiments with your cells.

 

   1. pipet a big drop of 0.1% poly-L lysine (PLL) in dH_2 O on your #
      1.5 cover slip
   2. allow the glass to coat for 30-60min at RT
   3. remove PLL as much as possible
   4. air dry till all liquid has evaporated
   5. dry for another hour at RT
   6. pipet drop of bead solution straight from the tube on the coated
      cover slip
   7. allow to adhere for 5 min
   8. pipet off as much as possible
   9. allow to dry for 30-60min in the dark
  10. put embedding medium on the microscope slide
  11. put cover slip with attached beads on the microscope slide
  12. allow curing embedding medium to cure
  13. seal cover slip to the slide with non-fluorescent nail-polish
  14. store at 4°C in the dark
  15. beads slides should remain re-usable for at least 6 months.



stu_the_flat wrote:

> Hi
>
> I'm new to this forum. I have tried searching this forum but I can't find a
> topic. I apologize if I missed it.
>
> I'm a PHD student I'm relatively new to confocal imaging I am trying to
> image sub resolution beads to measure the point spread function.
>
> I am using a bio rad system I use a 60X water immersion lens I set the
> system to 1024 X 1024 at 16 bit resolution. Kalman sampling was set to 4 and
> the scanning head was as slow as possible and the laser power was as low as
> possible to measure the beads.
>
> I have found that this is the perfect recipe for measuring photo bleaching!
>
> I tried looking around for some sort of protocol on imaging sub resolution
> beads but I was unable to find anything. Also as I understand it there will
> always be some element of photo bleaching? Is it easy to compensate for if I
> can calculate how much beaching as occurred.
>
> Similarly if anybody as any tips on creating the slides with the beads I
> would be grateful mine are quite messy.
>
> Many thanks
>
> Stuart McIntyre
>
>
>  

[Glen MacDonald-2]

Hi Stuart,
You've learned the first lesson of confocal microscopy!  The problem  
is you are probably following very outdated instructions for the  
confocal.  The manuals and factory training for older confocals  
emphasized noise reduction through slow scan rates and repetitive  
averaging.  These gave dim, noise-free images of bleached specimens.  
Start with the 'Normal' scan rate and Kalman averaging at 3.  This  
will scan with a 5-fold reduction in exposure and gives nearly the  
same noise reduction on our MRC-1024.  Normal scan is 2 us/pixel, Slow  
scan is 8 us with the resulting intensities divided by 4.

If you are deconvolving the PSF and your specimens, try collecting a  
stack at Normal rate, without averaging.

You can test noise reduction as a function of dwell time and averaging  
to see where the noise reduction levels off as a function of Kalman  
repetition to get a sense of how much averaging is really necessary  
for your system.

Apply a coverslip with immersion oil to a fluorescent plastic  
reference slide and focus about 20 microns into the plastic using your  
favorite oil lens, with a zoom of  4 to get a uniform intensity across  
the field.
  Adjust laser and Gain to fill the histogram about 3/4 for a single  
image using Normal scan speed and Direct mode. Use these settings for  
collecting images at Slow Scan and Normal Scan using the Direct mode  
and with a Kalman of 2 to 8 or so..
Move the slide to a new field for each capture.
Graph the mean and Std. dev for each resulting image. Then graph the  
coefficients of variance (var=Std. dev/mean intensity) for each image.

Regards,
Glen

On Nov 24, 2008, at 1:18 AM, stu_the_flat wrote:

> Hi
>
> I'm new to this forum. I have tried searching this forum but I can't  
> find a
> topic. I apologize if I missed it.
>
> I'm a PHD student I'm relatively new to confocal imaging I am trying  
> to
> image sub resolution beads to measure the point spread function.
>
> I am using a bio rad system I use a 60X water immersion lens I set the
> system to 1024 X 1024 at 16 bit resolution. Kalman sampling was set  
> to 4 and
> the scanning head was as slow as possible and the laser power was as  
> low as
> possible to measure the beads.
>
> I have found that this is the perfect recipe for measuring photo  
> bleaching!
>
> I tried looking around for some sort of protocol on imaging sub  
> resolution
> beads but I was unable to find anything. Also as I understand it  
> there will
> always be some element of photo bleaching? Is it easy to compensate  
> for if I
> can calculate how much beaching as occurred.
>
> Similarly if anybody as any tips on creating the slides with the  
> beads I
> would be grateful mine are quite messy.
>
> Many thanks
>
> Stuart McIntyre
>
>
> --
> View this message in context: http://n2.nabble.com/Protocol-for-imaging-micro-beads--tp1571444p1571444.html
> Sent from the Confocal Microscopy List mailing list archive at  
> Nabble.com.

[Ignatius, Mike]

If you don't want to fuss with mounting your own beads, Molecular Probes, now a part of Life Technologies, sells slides with 5 different sizes of beads already mounted.  

TetraSpeck(tm) Fluorescent Microspheres Size Kit (T14792):
Position 1: 4.0 μm TetraSpeck(tm) microspheres
Position 2: 1.0 μm TetraSpeck(tm) microspheres
Position 3: 0.5 μm TetraSpeck(tm) microspheres
Position 4: 0.2 μm TetraSpeck(tm) microspheres
Position 5: 0.1 μm TetraSpeck(tm) microspheres
Position 6: mounted mixture of all five sizes of TetraSpeck(tm) microspheres

Regards,

Mike Ignatius
Molecular Probes/Life Technologies  

-----Original Message-----
From: Confocal Microscopy List [mailto:[hidden email]] On Behalf Of stu_the_flat
Sent: Monday, November 24, 2008 1:18 AM
To: [hidden email]
Subject: Protocol for imaging micro beads?

Hi

I'm new to this forum. I have tried searching this forum but I can't find a
topic. I apologize if I missed it.

I'm a PHD student I'm relatively new to confocal imaging I am trying to
image sub resolution beads to measure the point spread function.

I am using a bio rad system I use a 60X water immersion lens I set the
system to 1024 X 1024 at 16 bit resolution. Kalman sampling was set to 4 and
the scanning head was as slow as possible and the laser power was as low as
possible to measure the beads.

I have found that this is the perfect recipe for measuring photo bleaching!

I tried looking around for some sort of protocol on imaging sub resolution
beads but I was unable to find anything. Also as I understand it there will
always be some element of photo bleaching? Is it easy to compensate for if I
can calculate how much beaching as occurred.

Similarly if anybody as any tips on creating the slides with the beads I
would be grateful mine are quite messy.

Many thanks

Stuart McIntyre


--
View this message in context: http://n2.nabble.com/Protocol-for-imaging-micro-beads--tp1571444p1571444.html
Sent from the Confocal Microscopy List mailing list archive at Nabble.com.

[Farid Jalali]

This is an excellent product but I have found the 0.2 and 0.1um beads to exhibit some degree of Brownian motion that I do not detect when I mount Invitrogen/Molecular Probes P-Speck beads myself using procedures similar to the ones provided in this thread. I also find the smaller Tetraspeck beads to be very faint compared to the 170nm P-Speck beads which are very bright. For what its worth, I really like the P-Speck beads for PSF determination and that they come in 4 different excitation/emission ranges is very convenient.

Cheers
Farid

2008/11/24 Ignatius, Mike <[hidden email]>

If you don't want to fuss with mounting your own beads, Molecular Probes, now a part of Life Technologies, sells slides with 5 different sizes of beads already mounted.

TetraSpeck(tm) Fluorescent Microspheres Size Kit (T14792):
Position 1: 4.0 μm TetraSpeck(tm) microspheres
Position 2: 1.0 μm TetraSpeck(tm) microspheres
Position 3: 0.5 μm TetraSpeck(tm) microspheres
Position 4: 0.2 μm TetraSpeck(tm) microspheres
Position 5: 0.1 μm TetraSpeck(tm) microspheres
Position 6: mounted mixture of all five sizes of TetraSpeck(tm) microspheres

Regards,

Mike Ignatius
Molecular Probes/Life Technologies

-----Original Message-----
From: Confocal Microscopy List [mailto:[hidden email]] On Behalf Of stu_the_flat
Sent: Monday, November 24, 2008 1:18 AM
To: [hidden email]
Subject: Protocol for imaging micro beads?

Hi

I'm new to this forum. I have tried searching this forum but I can't find a
topic. I apologize if I missed it.

I'm a PHD student I'm relatively new to confocal imaging I am trying to
image sub resolution beads to measure the point spread function.

I am using a bio rad system I use a 60X water immersion lens I set the
system to 1024 X 1024 at 16 bit resolution. Kalman sampling was set to 4 and
the scanning head was as slow as possible and the laser power was as low as
possible to measure the beads.

I have found that this is the perfect recipe for measuring photo bleaching!

I tried looking around for some sort of protocol on imaging sub resolution
beads but I was unable to find anything. Also as I understand it there will
always be some element of photo bleaching? Is it easy to compensate for if I
can calculate how much beaching as occurred.

Similarly if anybody as any tips on creating the slides with the beads I
would be grateful mine are quite messy.

Many thanks

Stuart McIntyre


--
View this message in context: http://n2.nabble.com/Protocol-for-imaging-micro-beads--tp1571444p1571444.html
Sent from the Confocal Microscopy List mailing list archive at Nabble.com.

--
Farid Jalali MSc
Program Leader- Cellular Imaging Core
Applied Molecular Oncology and Radiation Medicine Program
Princess Margaret Hospital (University  Health Network)
Toronto Medical Discovery Tower
Toronto, Canada
416-581-7754 STTARR at TMDT
416-581-7791 STTARR Microscopy Suite

[Sarah Kefayati]

Hi Stuart,

 

My whole MSc research I spent on imaging the beads. If you search through the forum you would see the questions that I asked and so many great answers that I got.

 

But in brief, I used .2um Tetraspeck beads. Spreading them on the slide let it dry and then putting the slide on the heater with the temp. close to body temp.( This helps in a way that when you pipette the water drop ,beads don't float in the water.). After couple minuets on the heater ,pouring the water drop puuting the coverslip and sealing with the nail polish.

 

If you want to measure the depth dependent PSF, great way would be making the sample with agarose gel. I used a metal filter frame( you can use whatever that you find handy ) .I put the frame on the coverslip ( using the bigger size of the coverslip), pouring 5um of the beads on the surface of the coverslip through the hole of the frame ( in this way you can identify 0um depth during the imaging). Afterwards, pipetting the hot gel liquid through the hole (~ .5 ml) and then pipetting about 20um of the beads on the center of the gel and mixing vertically in the center. let the gel solidify completely and then carefully detach your frame from the gel ( make sure that the gel is still attached to the coverslip while you are taking off the frame).

 

Hope it helps

let me know if you have any further questions

 

Sarah 

2008/11/24 Farid Jalali <[hidden email]>

This is an excellent product but I have found the 0.2 and 0.1um beads to exhibit some degree of Brownian motion that I do not detect when I mount Invitrogen/Molecular Probes P-Speck beads myself using procedures similar to the ones provided in this thread. I also find the smaller Tetraspeck beads to be very faint compared to the 170nm P-Speck beads which are very bright. For what its worth, I really like the P-Speck beads for PSF determination and that they come in 4 different excitation/emission ranges is very convenient.

Cheers
Farid

2008/11/24 Ignatius, Mike <[hidden email]>

If you don't want to fuss with mounting your own beads, Molecular Probes, now a part of Life Technologies, sells slides with 5 different sizes of beads already mounted.

TetraSpeck(tm) Fluorescent Microspheres Size Kit (T14792):
Position 1: 4.0 μm TetraSpeck(tm) microspheres
Position 2: 1.0 μm TetraSpeck(tm) microspheres
Position 3: 0.5 μm TetraSpeck(tm) microspheres
Position 4: 0.2 μm TetraSpeck(tm) microspheres
Position 5: 0.1 μm TetraSpeck(tm) microspheres
Position 6: mounted mixture of all five sizes of TetraSpeck(tm) microspheres

Regards,

Mike Ignatius
Molecular Probes/Life Technologies

-----Original Message-----
From: Confocal Microscopy List [mailto:[hidden email]] On Behalf Of stu_the_flat
Sent: Monday, November 24, 2008 1:18 AM
To: [hidden email]
Subject: Protocol for imaging micro beads?

Hi

I'm new to this forum. I have tried searching this forum but I can't find a
topic. I apologize if I missed it.

I'm a PHD student I'm relatively new to confocal imaging I am trying to
image sub resolution beads to measure the point spread function.

I am using a bio rad system I use a 60X water immersion lens I set the
system to 1024 X 1024 at 16 bit resolution. Kalman sampling was set to 4 and
the scanning head was as slow as possible and the laser power was as low as
possible to measure the beads.

I have found that this is the perfect recipe for measuring photo bleaching!

I tried looking around for some sort of protocol on imaging sub resolution
beads but I was unable to find anything. Also as I understand it there will
always be some element of photo bleaching? Is it easy to compensate for if I
can calculate how much beaching as occurred.

Similarly if anybody as any tips on creating the slides with the beads I
would be grateful mine are quite messy.

Many thanks

Stuart McIntyre


--
View this message in context: http://n2.nabble.com/Protocol-for-imaging-micro-beads--tp1571444p1571444.html
Sent from the Confocal Microscopy List mailing list archive at Nabble.com.

--
Farid Jalali MSc
Program Leader- Cellular Imaging Core
Applied Molecular Oncology and Radiation Medicine Program
Princess Margaret Hospital (University  Health Network)
Toronto Medical Discovery Tower
Toronto, Canada
416-581-7754 STTARR at TMDT
416-581-7791 STTARR Microscopy Suite

[stu_the_flat]

Wow! This is a fantastic response!

first of all I would like to say thank you to all of you would took the time to email me and reply to this thread.

I'm aware of the commercial products available however as I'm just starting my PhD and we have a small stalk pile of beads I think it would be better if I get my hands dirty learning some skills! I have one concern some of you recommend heating the plate so the beads adhere to them. How can you be sure that the beads are not becoming deformed and possibly broadening?

Another thing is I would quite like to get the PSF through the complete z axis. obviously a little invalid if they are adhering to the cover slip. I also think that it would be more representative of a biological sample suspended in the solution?

Another tactic I read was to mix the beads with non fluorescent beads (or beads that don't fluoresce at the lasers wavelength if you feeling rich) I thought this is  a fantastic idea as it doesn't matter if they stick. I was wondering if there are rejected beads in the manufacturing proses at could be bought cheaply for this purpose?

I had used 5.7 µm (mainly to make it easy to focus the z axis)  mixed with 0.049 µm beads. these where suspended in agar.

--------------------------------------------------------------------------------------------------

Hi Stuart,
 
What Z stepsize are you using, and what is the precision of you Z stage? (i.e., are you using a Z Galvo-type system, or just a regular Z stepping motor?)  What's the size of the beads that you are using?  In my experience, you don't necessarily need to average 4 frames, 2-3 is enough if you have a decent signal to noise ratio to start with. How much do you zoom in (i.e. what's your XY pixel size)?   I shouldn't think that using the slowest scan is advantageous either, I'd scan at more laser power and at a higher frequency.
  You'll always have some bleaching but the final result should be okay even without bleaching correction. You can buy pre-fabricated slides from Invitrogen/Molecular probes, they are great but expensive, still worth the money if you buy the multi-well version that has 4, 2, 1, 0.2, 0.1 um and mix. You can buy the suspension cheaper and then follow Mariette's protocol of course but then you'll end up with a  vial full of same-size beads that you'll hardly use unless you produce a new test slide very often.
 I hope this helps a bit,
 
Zoltan

-------------------------------------------------------------------------------------------------

I am using a humble Z step motor when imaging the 5.7 µm bead my z step was 0.5µm I then went to 0.2µm when imaging the 0.049 µm beads. I had the zoom set to the full 10X so I think my pixel size would represent 0.02 µm. I realise  that is ridiculously small I was simply trying to gain the highest image quality possible.

Once again thank you

Stuart McIntyre

[lgelman]

Hi Stuart,

 

In order to be able to assess and compare the performance of our different microscopes over time and also to be able to compare our PSFs with those obtained at other facilities, we have written a macro for ImageJ which I attach to this mail. We started also to distribute it to several facilities and hope we can collect after a while enough pictures to know really how microscopes in different facilities really perform.

 

We make Point Spread Functions every week for all objectives on our High-end microscopes on the facility. On our confocals, we typically take 256x256 pixel images, with a pixel size of 60 to 70nm (zoom is adjusted for each magnification). We actually make a stack with 100 planes separated by 200nm. Pixel dwell time ranges typically from 2us to 5us. Images are 12-bit.

 

In short, the Macro crops the image to make a region of 15uM around the center of the bead, which is determined by the user by right-clicking on it. It then makes XZ and XY projections of the PSF, displays them in a window of 550x550 pixels together with the MIP of the stack (I can send an example on request directly per e-mail). It also fit the PSF with a Gaussian function and extracts FWHM laterally and axially. The functions and the fits are displayed behind the PSF (one gets at the end a stack with 2 slices, the PSF on slice#1 and the curves on slice#2).

 

We use beads from Molecular probes (PS-Speck microscope point source kit, P-7220) and never experienced any photobleaching. We make a 1/10’000 dilution of the beads, lay tiny drops on a slide and let them dry. Then we add a drop mounting medium (in our case Prolong Gold) and cover with a coverslide.

 

Do not hesitate to contact me if you have any question or idea to improve the Macro. I would be glad also if you would accept after a while to send me some of your PSFs (even if you don’t work on a facility), so I could compare all the PSFs I would get from different places. I would send this back to all interested users, so everybody can know how his scopes perform in comparison with others.

 

Very best regards,

 

Laurent.

 

 

 

________________

 

Installation and use of the Macro:

 

You need to install the each time you start ImageJ. Go to >Plugins>Macros>Install… Select it in the dialog window and click "open".

 

To run it, you need before to open a stack. Remember, we take stacks of 100 planes, spaced by 0.2um, for all objectives and all microscopes.

Then go to >Plugins>Macros>MIPs for PSFs for all microscopes V2 to run the Macro.

 

Automatic Macro actions / User actions:

A. Selects the plane with the highest pixel intensity, adjusts display settings, opens the information dialog box.

1. Enter information in the dialog window which popped up.

2. Zoom in the image to clearly localize the center of the bead (you can also navigate between planes if needed).

3. Right clicks with the mouse on the center of the bead.

B. Crops the image to get 15umx15um area centered over the pixel clicked by the user.

C. Makes projections in X and Y of the stack

D. Stitches together the cropped area and the projections

E. Estimates and subtracts background

F. Takes the square root of the image (to minimize photon noise and to mimic a decrease in histogram gain)

G. Resizes the image to 550x550 pixels, adjusts display, changes LUT and rename the picture with a standardized name: Date_Scope name_Magnification_NA.

H. Extracts FWHM and displays curves.

 

Of course you can customize the Macro as we did, for example we don't enter the name of the scope but we have a scrolling list with our scopes and the pixel size is then calculated automatically based on the magnification and the pixel size of our camera chips (for wide-field microscopes). Only when a LSM scope is selected a second window pops up asking for pixel size. We have also an additional information about the presence of an optovar in some of our systems.

 

 

______________________________________

Laurent Gelman, PhD

Friedrich Miescher Institut

Facility for Advanced Imaging and Microscopy

WRO 1066.2.16

Maulbeerstrasse 66

CH-4058 Basel

Tel.: [phone] / Cell phone: [phone]

www.fmi.ch

[hidden email]

 

 

 

[lgelman]

 

From: Gelman, Laurent
Sent: mardi, 25. novembre 2008 11:20
To: 'Confocal Microscopy List'
Subject: RE: Protocol for imaging micro beads?

 

Hi Stuart,

 

In order to be able to assess and compare the performance of our different microscopes over time and also to be able to compare our PSFs with those obtained at other facilities, we have written a macro for ImageJ which I attach to this mail. We started also to distribute it to several facilities and hope we can collect after a while enough pictures to know really how microscopes in different facilities really perform.

 

We make Point Spread Functions every week for all objectives on our High-end microscopes on the facility. On our confocals, we typically take 256x256 pixel images, with a pixel size of 60 to 70nm (zoom is adjusted for each magnification). We actually make a stack with 100 planes separated by 200nm. Pixel dwell time ranges typically from 2us to 5us. Images are 12-bit.

 

In short, the Macro crops the image to make a region of 15uM around the center of the bead, which is determined by the user by right-clicking on it. It then makes XZ and XY projections of the PSF, displays them in a window of 550x550 pixels together with the MIP of the stack (I can send an example on request directly per e-mail). It also fit the PSF with a Gaussian function and extracts FWHM laterally and axially. The functions and the fits are displayed behind the PSF (one gets at the end a stack with 2 slices, the PSF on slice#1 and the curves on slice#2).

 

We use beads from Molecular probes (PS-Speck microscope point source kit, P-7220) and never experienced any photobleaching. We make a 1/10’000 dilution of the beads, lay tiny drops on a slide and let them dry. Then we add a drop mounting medium (in our case Prolong Gold) and cover with a coverslide.

 

Do not hesitate to contact me if you have any question or idea to improve the Macro. I would be glad also if you would accept after a while to send me some of your PSFs (even if you don’t work on a facility), so I could compare all the PSFs I would get from different places. I would send this back to all interested users, so everybody can know how his scopes perform in comparison with others.

 

Very best regards,

 

Laurent.

 

 

 

________________

 

Installation and use of the Macro:

 

You need to install the each time you start ImageJ. Go to >Plugins>Macros>Install… Select it in the dialog window and click "open".

 

To run it, you need before to open a stack. Remember, we take stacks of 100 planes, spaced by 0.2um, for all objectives and all microscopes.

Then go to >Plugins>Macros>MIPs for PSFs for all microscopes V2 to run the Macro.

 

Automatic Macro actions / User actions:

A. Selects the plane with the highest pixel intensity, adjusts display settings, opens the information dialog box.

1. Enter information in the dialog window which popped up.

2. Zoom in the image to clearly localize the center of the bead (you can also navigate between planes if needed).

3. Right clicks with the mouse on the center of the bead.

B. Crops the image to get 15umx15um area centered over the pixel clicked by the user.

C. Makes projections in X and Y of the stack

D. Stitches together the cropped area and the projections

E. Estimates and subtracts background

F. Takes the square root of the image (to minimize photon noise and to mimic a decrease in histogram gain)

G. Resizes the image to 550x550 pixels, adjusts display, changes LUT and rename the picture with a standardized name: Date_Scope name_Magnification_NA.

H. Extracts FWHM and displays curves.

 

Of course you can customize the Macro as we did, for example we don't enter the name of the scope but we have a scrolling list with our scopes and the pixel size is then calculated automatically based on the magnification and the pixel size of our camera chips (for wide-field microscopes). Only when a LSM scope is selected a second window pops up asking for pixel size. We have also an additional information about the presence of an optovar in some of our systems.

 

 

______________________________________

Laurent Gelman, PhD

Friedrich Miescher Institut

Facility for Advanced Imaging and Microscopy

WRO 1066.2.16

Maulbeerstrasse 66

CH-4058 Basel

Tel.: [phone] / Cell phone: [phone]

www.fmi.ch

[hidden email]

 

 

 

[Glen MacDonald-2]

Dear Stuart,
Why are you using .049 um beads? A PSF bead only needs to be smaller  
than the optical resolution of the objective lens. compare the volume  
of that bead to a 120 or 170 um bead to get an idea of the difference  
in possible number of fluorophores available to generate a signal.  
Such small beads and prolonged scanning creates an exercise in  
photobleaching. The RI difference between coverglass and the bead can  
definitely be an issue. However, if you are trying to generate a PSF  
for imaging monolayers of cells adhered to glass, that could be a  
useful PSF. A mounting medium with an RI close to that of the glass  
would reduce the coverglass issue.  A bigger issue when working close  
to the coverslip may be reflection.  The question is why are  
generating the PSF? To characterize the microscope or to describe the  
imaging conditions through a sample? I find that adding a few larger  
beads, ca 6 um or 15 um, is very useful for finding the coverslip.  
they also create a spacer for when I mount the beads. For a thicker  
volume, DAKO Glycergel (RI 1.44-1.47) or Fluormount G (RI 1.39) are  
useful hardening media. I've heard reports of latex beads melting in  
agarose that is too hot.

Regards,
Glen

> Wow! This is a fantastic response!
>
> first of all I would like to say thank you to all of you would took  
> the time
> to email me and reply to this thread.
>
> I'm aware of the commercial products available however as I'm just  
> starting
> my PhD and we have a small stalk pile of beads I think it would be  
> better if
> I get my hands dirty learning some skills! I have one concern some  
> of you
> recommend heating the plate so the beads adhere to them. How can you  
> be sure
> that the beads are not becoming deformed and possibly broadening?
>
> Another thing is I would quite like to get the PSF through the  
> complete z
> axis. obviously a little invalid if they are adhering to the cover  
> slip. I
> also think that it would be more representative of a biological sample
> suspended in the solution?
>
> Another tactic I read was to mix the beads with non fluorescent  
> beads (or
> beads that don't fluoresce at the lasers wavelength if you feeling  
> rich) I
> thought this is  a fantastic idea as it doesn't matter if they  
> stick. I was
> wondering if there are rejected beads in the manufacturing proses at  
> could
> be bought cheaply for this purpose?
>
> I had used 5.7 µm (mainly to make it easy to focus the z axis)  
> mixed with
> 0.049 µm beads. these where suspended in agar.
>
> --------------------------------------------------------------------------------------------------
>
> Hi Stuart,
>
> What Z stepsize are you using, and what is the precision of you Z  
> stage?
> (i.e., are you using a Z Galvo-type system, or just a regular Z  
> stepping
> motor?)  What's the size of the beads that you are using?  In my  
> experience,
> you don't necessarily need to average 4 frames, 2-3 is enough if you  
> have a
> decent signal to noise ratio to start with. How much do you zoom in  
> (i.e.
> what's your XY pixel size)?   I shouldn't think that using the  
> slowest scan
> is advantageous either, I'd scan at more laser power and at a higher
> frequency.
>  You'll always have some bleaching but the final result should be  
> okay even
> without bleaching correction. You can buy pre-fabricated slides from
> Invitrogen/Molecular probes, they are great but expensive, still  
> worth the
> money if you buy the multi-well version that has 4, 2, 1, 0.2, 0.1  
> um and
> mix. You can buy the suspension cheaper and then follow Mariette's  
> protocol
> of course but then you'll end up with a  vial full of same-size  
> beads that
> you'll hardly use unless you produce a new test slide very often.
> I hope this helps a bit,
>
> Zoltan
>
> -------------------------------------------------------------------------------------------------
>
> I am using a humble Z step motor when imaging the 5.7 µm bead my z  
> step was
> 0.5µm I then went to 0.2µm when imaging the 0.049 µm beads. I had  
> the zoom
> set to the full 10X so I think my pixel size would represent 0.02  
> µm. I
> realise  that is ridiculously small I was simply trying to gain the  
> highest
> image quality possible.
>
> Once again thank you
>
> Stuart McIntyre
> --
> View this message in context: http://n2.nabble.com/Protocol-for-imaging-micro-beads--tp1571444p1575704.html
> Sent from the Confocal Microscopy List mailing list archive at  
> Nabble.com.

[Sam's Mail]

Anyone have some good suggestions for a user who is having some challenges getting complete Ab penetration into her 50 micron thick brain slices (frozen).  It appears that her Ab's are only penetrating about 15 microns or so.  I believe she is using a reasonably standard IF protocol of 4% formaldehyde and 0.1% Triton X-100.

Thanks all for the ideas,

--
Samuel A. Connell
Director of Light Microscopy
Cell & Tissue Imaging Center
St. Jude Children's Research Hospital
262 Danny Thomas Place, D1052A
Memphis, TN 38105-3678
CTIC [phone]
Office [phone]
Cell [phone]
[hidden email]



________________________________
Email Disclaimer: www.stjude.org/emaildisclaimer

[anh2006]

Up the Triton to 0.3%.
Block in that + serum for 8+ hours
Do Ab incubations for 36-48 hours (in the blocking buffer).

>Anyone have some good suggestions for a user who is having some
>challenges getting complete Ab penetration into her 50 micron thick
>brain slices (frozen).  It appears that her Ab's are only
>penetrating about 15 microns or so.  I believe she is using a
>reasonably standard IF protocol of 4% formaldehyde and 0.1% Triton
>X-100.
>
>Thanks all for the ideas,
>
>--
>Samuel A. Connell
>Director of Light Microscopy
>Cell & Tissue Imaging Center
>St. Jude Children's Research Hospital
>262 Danny Thomas Place, D1052A
>Memphis, TN 38105-3678
>CTIC [phone]
>Office [phone]
>Cell [phone]
>[hidden email]
>
>
>
>________________________________
>Email Disclaimer: www.stjude.org/emaildisclaimer

--

[Glen MacDonald-2]

How long are her incubations?  Are these on slides or free-floating?
Most people make the mistake of going overnight or longer in primary  
then only 1-2 hours in secondary.  Both should incubate at least  
overnight.  The need for Triton should be tested, it may  be unnecessary

Regards,
Glen.

Glen MacDonald
Core for Communication Research
Virginia Merrill Bloedel Hearing Research Center
Box 357923
University of Washington
Seattle, WA 98195-7923  USA
[phone]
[hidden email]

******************************************************************************
The box said "Requires WindowsXP or better", so I bought a Macintosh.
******************************************************************************


On Jan 9, 2009, at 5:06 PM, Connell, Samuel wrote:

> Anyone have some good suggestions for a user who is having some  
> challenges getting complete Ab penetration into her 50 micron thick  
> brain slices (frozen).  It appears that her Ab's are only  
> penetrating about 15 microns or so.  I believe she is using a  
> reasonably standard IF protocol of 4% formaldehyde and 0.1% Triton  
> X-100.
>
> Thanks all for the ideas,
>
> --
> Samuel A. Connell
> Director of Light Microscopy
> Cell & Tissue Imaging Center
> St. Jude Children's Research Hospital
> 262 Danny Thomas Place, D1052A
> Memphis, TN 38105-3678
> CTIC [phone]
> Office [phone]
> Cell [phone]
> [hidden email]
>
>
>
> ________________________________
> Email Disclaimer: www.stjude.org/emaildisclaimer

[Stephen C. Kempf]

Dehydrate sections to 70% ethanol with a dehydration series (30%, 50%,  
70% EtOH) and then rehydrate to PBS and proceed with labeling. For 50  
um sections, 5 - 10 min per step should be fine. This method has  
worked very well for us on larvae that are 80 - 300 um in diameter.  
It's possible that dehydration to 30% or 50% EtOH might work, but  
you'd have to try it and see.

Steve Kempf
Associate Professor
Faculty Director, AU Hybridoma Facility
331 Funchess Hall
Dept. of Biological Sciences
Auburn University
Auburn, AL  36849
Tel: [phone]

On Jan 9, 2009, at 7:06 PM, Connell, Samuel wrote:

> Anyone have some good suggestions for a user who is having some  
> challenges getting complete Ab penetration into her 50 micron thick  
> brain slices (frozen).  It appears that her Ab's are only  
> penetrating about 15 microns or so.  I believe she is using a  
> reasonably standard IF protocol of 4% formaldehyde and 0.1% Triton  
> X-100.
>
> Thanks all for the ideas,
>
> --
> Samuel A. Connell
> Director of Light Microscopy
> Cell & Tissue Imaging Center
> St. Jude Children's Research Hospital
> 262 Danny Thomas Place, D1052A
> Memphis, TN 38105-3678
> CTIC [phone]
> Office [phone]
> Cell [phone]
> [hidden email]
>
>
>
> ________________________________
> Email Disclaimer: www.stjude.org/emaildisclaimer

[RICHARD BURRY]

Samuel
 
In my experience, the key issue in penetration is incubation temperature.  4oC is too cold, Triton does not work well at the 4oC.  Remaining lipids at 4 oC change phase further inhibiting penetration.  Room temperature for 24 or 48 hrs is best.  Besure to use 0.02% sodium azide in the buffer to kill any bacteria. 
 
Dick

----- Original Message -----
From: "Connell, Samuel" <[hidden email]>
Date: Friday, January 9, 2009 8:05 pm
Subject: Ab Penetration
To: [hidden email]

> Anyone have some good suggestions for a user who is having some
> challenges getting complete Ab penetration into her 50 micron
> thick brain slices (frozen).  It appears that her Ab's are
> only penetrating about 15 microns or so.  I believe she is
> using a reasonably standard IF protocol of 4% formaldehyde and
> 0.1% Triton X-100.
>
> Thanks all for the ideas,
>
> --
> Samuel A. Connell
> Director of Light Microscopy
> Cell & Tissue Imaging Center
> St. Jude Children's Research Hospital
> 262 Danny Thomas Place, D1052A
> Memphis, TN 38105-3678
> CTIC [phone]
> Office [phone]
> Cell [phone]
> [hidden email]
>
>
>
> ________________________________
> Email Disclaimer: www.stjude.org/emaildisclaimer
>
>
> --
> BEGIN-ANTISPAM-VOTING-LINKS
> ------------------------------------------------------
>
> Teach CanIt if this mail (ID 778805836) is spam:
> Spam:       
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Richard W. Burry, Ph.D.
Department of Neuroscience, College of Medicine
Campus Microscopy and Imaging Facility, Director
The Ohio State University
Associate Editor, Journal of Histochemistry and Cytochemistry
277 Biomedical Research Tower
460 West Twelfth Avenue
Columbus, Ohio 43210
Voice 614.292.2814  Cell 614.638.3345  Fax [phone]

[Mollie Lange]

Use free-floating sections if you can.  Agitate on a gyrotary mixer during all antibody and rinse steps.  Leave in primary antibody for at least two nights at 4 degrees C on the shaker (we use the coldroom). The secondary antibody incubation can be extended also. These longer incubations will give the triton time to do its work and time for the antibodies to penetrate. It the tissue can take it, increase the triton to .15 or .2%.

Mollie Lange
Intl Ctr for Spinal Cord Injury
Kennedy Krieger Institute

>>> "Connell, Samuel" <[hidden email]> 1/9/2009 8:06 PM >>>
Anyone have some good suggestions for a user who is having some challenges getting complete Ab penetration into her 50 micron thick brain slices (frozen).  It appears that her Ab's are only penetrating about 15 microns or so.  I believe she is using a reasonably standard IF protocol of 4% formaldehyde and 0.1% Triton X-100.

Thanks all for the ideas,

--
Samuel A. Connell
Director of Light Microscopy
Cell & Tissue Imaging Center
St. Jude Children's Research Hospital
262 Danny Thomas Place, D1052A
Memphis, TN 38105-3678
CTIC [phone]
Office [phone]
Cell [phone]
[hidden email]



________________________________
Email Disclaimer: www.stjude.org/emaildisclaimer


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[Nathan-64]

Hi all,

Sorry to be a bit late on this, but I've used the following CSHL protocol for tagging various proteins in rat (SD) eye cup whole-mount preparations:
 
  http://cshprotocols.cshlp.org/cgi/content/full/2008/3/pdb.prot4957

Best of luck,
Nate


--
Nathan O'Connor
Silver Laboratory
Physiology and Biophysics
Weill Cornell Medical College
New York, NY 10021

On Sun, Jan 11, 2009 at 7:42 PM, Mollie Lange <[hidden email]> wrote:

Use free-floating sections if you can.  Agitate on a gyrotary mixer during all antibody and rinse steps.  Leave in primary antibody for at least two nights at 4 degrees C on the shaker (we use the coldroom). The secondary antibody incubation can be extended also. These longer incubations will give the triton time to do its work and time for the antibodies to penetrate. It the tissue can take it, increase the triton to .15 or .2%.

Mollie Lange
Intl Ctr for Spinal Cord Injury
Kennedy Krieger Institute

>>> "Connell, Samuel" <[hidden email]> 1/9/2009 8:06 PM >>>

Anyone have some good suggestions for a user who is having some challenges getting complete Ab penetration into her 50 micron thick brain slices (frozen).  It appears that her Ab's are only penetrating about 15 microns or so.  I believe she is using a reasonably standard IF protocol of 4% formaldehyde and 0.1% Triton X-100.

Thanks all for the ideas,

--
Samuel A. Connell
Director of Light Microscopy
Cell & Tissue Imaging Center
St. Jude Children's Research Hospital
262 Danny Thomas Place, D1052A
Memphis, TN 38105-3678
CTIC [phone]
Office [phone]
Cell [phone]
[hidden email]



________________________________
Email Disclaimer: www.stjude.org/emaildisclaimer

Please consider the environment before printing this E-Mail.



Disclaimer:
The materials in this e-mail are private and may contain Protected Health Information. Please note that e-mail is not necessarily confidential or secure. Your use of e-mail constitutes your acknowledgment of these confidentiality and security limitations. If you are not the intended recipient, be advised that any unauthorized use, disclosure, copying, distribution, or the taking of any action in reliance on the contents of this information is strictly prohibited. If you have received this e-mail in error, please immediately notify the sender via telephone or return e-mail.