Re: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes of Kidney - Issues and Questions

[Colleen Forster via Histonet]

Make sure the periodic acid is made fresh EACH time you run  the stain.
That can also make a big difference in the stain quality.

Colleen Forster HT(ASCP)QIHC

On Thu, Sep 23, 2021 at 6:14 PM Tony Henwood (SCHN) via Histonet <
histonet@lists.utsouthwestern.edu> wrote:

> I agree with Bryan,
>
> The introduction of thiosemicarbazide before the silver step improves the
> staining immensely.
>
> I would also look at the periodic acid. Is it too dilute, though 0.5%
> should work? I usually cover this by using a 1% solution for 20 minutes.
>
> Regards
> Tony Henwood JP, MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA)
> Principal Scientist, the Children’s Hospital at Westmead
> Adjunct Fellow, School of Medicine, University of Western Sydney
> Tel: 612 9845 3306
> Fax: 612 9845 3318
> Pathology Department
> the children's hospital at westmead
> Cnr Hawkesbury Road and Hainsworth Street, Westmead
> Locked Bag 4001, Westmead NSW 2145, AUSTRALIA
>
>
> -Original Message-
> From: Bryan Llewellyn via Histonet [mailto:
> histonet@lists.utsouthwestern.edu]
> Sent: Friday, 24 September 2021 7:47 AM
> To: Jordan ; Histonet <
> histonet@lists.utsouthwestern.edu>
> Subject: Re: [Histonet] Jones' Methenamine Silver Stain for Basement
> Membranes of Kidney - Issues and Questions
>
> Hi,
> Try the method given in StainsFile at:
> http://stainsfile.info/stain/metallic/jones.htm
>
> Bryan Llewellyn
>
>
> Hood, Jordan via Histonet wrote:
> > Hello,
> >
> > I'm new to histology (and new to histonet), and I work in a small
> histology lab specializing in animal tissues that receives
> requests/submissions from researchers. I tried (and failed) to perform a
> Jones' Methenamine Silver stain on a client's submission of pig kidneys
> (formalin-fixed, paraffin-embedded, cut at 2.5 microns), and I need some
> help troubleshooting this stain since my co-workers are stumped, too.  I
> used the following procedure from Rowley Biochemical:
> >
> >
> > ~
> > "Fixation: 10% Buffered Neutral Formalin (F-113) or Bouin's Solution
> > (F-40) or Zenker's (F-155)
> >
> > Sections: Paraffin, 2 microns
> >
> > Procedure: Acid washed glassware must be used
> > 1. Deparaffinize and hydrate to distilled water.
> > 2. Oxidize in Periodic Acid 0.5% (F-396-1) 11 minutes. Wash in
> chloride-free water.
> > 3. Prepare Methenamine Silver solution by mixing: 42.5 ml Methenamine 3%
> (F-396-2), 2.5 ml Silver Nitrate, 5% (F-396-3) and 12.0 ml Borate Buffer,
> pH 8.2 (F-396-4).
> > 4. Place slides in the solution and the entire jar in a water bath at
> 70°C for approx. 60-75 minutes. Check under microscope when slides appear
> medium brown microscopically. Every 10 minutes, once the medium brown color
> has been established, rinse a slide in 70°C, chloride free water and check
> under a microscope. Rinse again in hot water and return to the hot staining
> solution. As the staining time approaches the end point, check the slides,
> as above, every 1-2 minutes. The entire procedure must be performed quickly
> to prevent an uneven staining of the tissues. The slides should exhibit a
> brownish- yellow background, intense black reticulum fibers, and black
> basement membranes. If the slides become oversaturated, i.e. too black,
> destain in a dilute Potassium Ferricyanide Solution (F-396-11) for one or
> two dips.
> > 5. Rinse well in distilled water. Tone in Gold Chloride 0.2% (F-396-5),
> 1 minute. If sections are overtoned place in Sodium Metabisulfite, 3%
> (F-396-12) for 1-3 minutes. Rinse well in distilled water.
> > 6. Sodium Thiosulfate 3% (F-396-9), 1-2 miutes. Wash in running tap
> water, 10 minutes. Rinse well in distilled water.
> > 7. Stain in Harris' Hematoxylin (F-396-6) containing 2-4ml of Glacial
> Acetic Acid per 100 ml for 5-15 minutes. Wash in water.
> > 8. Differentiate in Acid Alcohol 1% (F-396-13) until the sections turn
> red.
> > 9. Blue section in Ammonia Water, 0.3% (F-396-14). Wash thoroughly.
> > 10. Counterstain in Eosin Y, 1%, Alcoholic Solution (F-396-7).
> > 11. Dehydrate in 95% alcohol, absolute alcohol and clear in xylene 3
> changes each. Mount.
> >
> > Stain Results:
> > Basement membranes, reticulum fibers: Black
> > Nuclei: Blue
> > Cytoplasm, collagen, connective tissue: Pink-orange
> >
> > References: Jones, D.B., Amer.J.Path. 27:99 (1951). AFIP Manual of
> Histolocical Staining Methods, 3rd ed., Ed. L. Luna: NY: McGraw-Hill
> Publications, c. 1968, p. 97."
> > ~
> >
> >
> > It became apparent that something went wrong during Step 4 when the
> slides were in the glass container (not a coplin jar - we have ten slides
> that we need to stain so we're using a rectangular glass container that
> holds ten slides on their sides - it does require a metal handle to move,
> but the handle is flexible and easy to remove after the glass slide rack
> has been transferred between containers) of silver solution in the water
> bath because there was lots of precipitate on the slides and floating on
> the surface of th

Re: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes of Kidney - Issues and Questions

[Tony Henwood (SCHN) via Histonet]

I agree with Bryan,

The introduction of thiosemicarbazide before the silver step improves the 
staining immensely.

I would also look at the periodic acid. Is it too dilute, though 0.5% should 
work? I usually cover this by using a 1% solution for 20 minutes.

Regards 
Tony Henwood JP, MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA) 
Principal Scientist, the Children’s Hospital at Westmead
Adjunct Fellow, School of Medicine, University of Western Sydney 
Tel: 612 9845 3306 
Fax: 612 9845 3318 
Pathology Department
the children's hospital at westmead
Cnr Hawkesbury Road and Hainsworth Street, Westmead
Locked Bag 4001, Westmead NSW 2145, AUSTRALIA 


-Original Message-
From: Bryan Llewellyn via Histonet [mailto:histonet@lists.utsouthwestern.edu] 
Sent: Friday, 24 September 2021 7:47 AM
To: Jordan ; Histonet 

Subject: Re: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes 
of Kidney - Issues and Questions

Hi,
Try the method given in StainsFile at:
http://stainsfile.info/stain/metallic/jones.htm

Bryan Llewellyn


Hood, Jordan via Histonet wrote:
> Hello,
> 
> I'm new to histology (and new to histonet), and I work in a small histology 
> lab specializing in animal tissues that receives requests/submissions from 
> researchers. I tried (and failed) to perform a Jones' Methenamine Silver 
> stain on a client's submission of pig kidneys (formalin-fixed, 
> paraffin-embedded, cut at 2.5 microns), and I need some help troubleshooting 
> this stain since my co-workers are stumped, too.  I used the following 
> procedure from Rowley Biochemical:
> 
> 
> ~
> "Fixation: 10% Buffered Neutral Formalin (F-113) or Bouin's Solution 
> (F-40) or Zenker's (F-155)
> 
> Sections: Paraffin, 2 microns
> 
> Procedure: Acid washed glassware must be used
> 1. Deparaffinize and hydrate to distilled water.
> 2. Oxidize in Periodic Acid 0.5% (F-396-1) 11 minutes. Wash in chloride-free 
> water.
> 3. Prepare Methenamine Silver solution by mixing: 42.5 ml Methenamine 3% 
> (F-396-2), 2.5 ml Silver Nitrate, 5% (F-396-3) and 12.0 ml Borate Buffer, pH 
> 8.2 (F-396-4).
> 4. Place slides in the solution and the entire jar in a water bath at 70°C 
> for approx. 60-75 minutes. Check under microscope when slides appear medium 
> brown microscopically. Every 10 minutes, once the medium brown color has been 
> established, rinse a slide in 70°C, chloride free water and check under a 
> microscope. Rinse again in hot water and return to the hot staining solution. 
> As the staining time approaches the end point, check the slides, as above, 
> every 1-2 minutes. The entire procedure must be performed quickly to prevent 
> an uneven staining of the tissues. The slides should exhibit a brownish- 
> yellow background, intense black reticulum fibers, and black basement 
> membranes. If the slides become oversaturated, i.e. too black, destain in a 
> dilute Potassium Ferricyanide Solution (F-396-11) for one or two dips.
> 5. Rinse well in distilled water. Tone in Gold Chloride 0.2% (F-396-5), 1 
> minute. If sections are overtoned place in Sodium Metabisulfite, 3% 
> (F-396-12) for 1-3 minutes. Rinse well in distilled water.
> 6. Sodium Thiosulfate 3% (F-396-9), 1-2 miutes. Wash in running tap water, 10 
> minutes. Rinse well in distilled water.
> 7. Stain in Harris' Hematoxylin (F-396-6) containing 2-4ml of Glacial Acetic 
> Acid per 100 ml for 5-15 minutes. Wash in water.
> 8. Differentiate in Acid Alcohol 1% (F-396-13) until the sections turn red.
> 9. Blue section in Ammonia Water, 0.3% (F-396-14). Wash thoroughly.
> 10. Counterstain in Eosin Y, 1%, Alcoholic Solution (F-396-7).
> 11. Dehydrate in 95% alcohol, absolute alcohol and clear in xylene 3 changes 
> each. Mount.
> 
> Stain Results:
> Basement membranes, reticulum fibers: Black
> Nuclei: Blue
> Cytoplasm, collagen, connective tissue: Pink-orange
> 
> References: Jones, D.B., Amer.J.Path. 27:99 (1951). AFIP Manual of 
> Histolocical Staining Methods, 3rd ed., Ed. L. Luna: NY: McGraw-Hill 
> Publications, c. 1968, p. 97."
> ~
> 
> 
> It became apparent that something went wrong during Step 4 when the slides 
> were in the glass container (not a coplin jar - we have ten slides that we 
> need to stain so we're using a rectangular glass container that holds ten 
> slides on their sides - it does require a metal handle to move, but the 
> handle is flexible and easy to remove after the glass slide rack has been 
> transferred between containers) of silver solution in the water bath because 
> there was lots of precipitate on the slides and floating on the surface of 
> the silver solution.
> 
> In my first test, I used five test slides (extra slides that we cut from the 
> same blocks that were submitted to us). I deparaffinized them in coplin jars 
> (moving them with plastic forceps) and hydrated them to deionized water. I 
> transferred the slides to a glass slide rack that holds ten slides on their 
> sides, added five blank slides that were rinsed in

Re: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes of Kidney - Issues and Questions

[Bryan Llewellyn via Histonet]

Hi,
Try the method given in StainsFile at:
http://stainsfile.info/stain/metallic/jones.htm

Bryan Llewellyn


Hood, Jordan via Histonet wrote:

Hello,

I'm new to histology (and new to histonet), and I work in a small histology lab 
specializing in animal tissues that receives requests/submissions from 
researchers. I tried (and failed) to perform a Jones' Methenamine Silver stain 
on a client's submission of pig kidneys (formalin-fixed, paraffin-embedded, cut 
at 2.5 microns), and I need some help troubleshooting this stain since my 
co-workers are stumped, too.  I used the following procedure from Rowley 
Biochemical:


~
"Fixation: 10% Buffered Neutral Formalin (F-113) or Bouin's Solution (F-40) or 
Zenker's (F-155)

Sections: Paraffin, 2 microns

Procedure: Acid washed glassware must be used
1. Deparaffinize and hydrate to distilled water.
2. Oxidize in Periodic Acid 0.5% (F-396-1) 11 minutes. Wash in chloride-free 
water.
3. Prepare Methenamine Silver solution by mixing: 42.5 ml Methenamine 3% 
(F-396-2), 2.5 ml Silver Nitrate, 5% (F-396-3) and 12.0 ml Borate Buffer, pH 
8.2 (F-396-4).
4. Place slides in the solution and the entire jar in a water bath at 70°C for 
approx. 60-75 minutes. Check under microscope when slides appear medium brown 
microscopically. Every 10 minutes, once the medium brown color has been 
established, rinse a slide in 70°C, chloride free water and check under a 
microscope. Rinse again in hot water and return to the hot staining solution. 
As the staining time approaches the end point, check the slides, as above, 
every 1-2 minutes. The entire procedure must be performed quickly to prevent an 
uneven staining of the tissues. The slides should exhibit a brownish- yellow 
background, intense black reticulum fibers, and black basement membranes. If 
the slides become oversaturated, i.e. too black, destain in a dilute Potassium 
Ferricyanide Solution (F-396-11) for one or two dips.
5. Rinse well in distilled water. Tone in Gold Chloride 0.2% (F-396-5), 1 
minute. If sections are overtoned place in Sodium Metabisulfite, 3% (F-396-12) 
for 1-3 minutes. Rinse well in distilled water.
6. Sodium Thiosulfate 3% (F-396-9), 1-2 miutes. Wash in running tap water, 10 
minutes. Rinse well in distilled water.
7. Stain in Harris' Hematoxylin (F-396-6) containing 2-4ml of Glacial Acetic 
Acid per 100 ml for 5-15 minutes. Wash in water.
8. Differentiate in Acid Alcohol 1% (F-396-13) until the sections turn red.
9. Blue section in Ammonia Water, 0.3% (F-396-14). Wash thoroughly.
10. Counterstain in Eosin Y, 1%, Alcoholic Solution (F-396-7).
11. Dehydrate in 95% alcohol, absolute alcohol and clear in xylene 3 changes 
each. Mount.

Stain Results:
Basement membranes, reticulum fibers: Black
Nuclei: Blue
Cytoplasm, collagen, connective tissue: Pink-orange

References: Jones, D.B., Amer.J.Path. 27:99 (1951). AFIP Manual of Histolocical 
Staining Methods, 3rd ed., Ed. L. Luna: NY: McGraw-Hill Publications, c. 1968, p. 
97."
~


It became apparent that something went wrong during Step 4 when the slides were 
in the glass container (not a coplin jar - we have ten slides that we need to 
stain so we're using a rectangular glass container that holds ten slides on 
their sides - it does require a metal handle to move, but the handle is 
flexible and easy to remove after the glass slide rack has been transferred 
between containers) of silver solution in the water bath because there was lots 
of precipitate on the slides and floating on the surface of the silver solution.

In my first test, I used five test slides (extra slides that we cut from the same blocks 
that were submitted to us). I deparaffinized them in coplin jars (moving them with 
plastic forceps) and hydrated them to deionized water. I transferred the slides to a 
glass slide rack that holds ten slides on their sides, added five blank slides that were 
rinsed in deionized water (so that the displacement of reagents would be equivalent to 
when we stain our ten "real" slides after testing is complete), and completed 
Step 2. I don't recall exactly how long the glass container of silver solution and the 
glass container of deionized water had been heating up in the water bath, but I would 
estimate ~15-30 minutes. The thermometer said that the water in the bath (not inside the 
containers) reached ~60-65 degrees Celsius. The silver solution was clear and colorless 
when I made it up, but by the time I put the slides into the warm silver solution, the 
solution was beginning to turn a light brown color (though it was still clear and I did 
not see any precipitate floating around). I removed the metal handle of the glass slide 
rack after the rack was transferred into the silver solution, but the metal handle did 
dip into the silver solution briefly. At some point, I noticed precipitate floating 
around of the surface of the silver solution. After ~80 minutes, I used plastic forceps 
to remove one test slide from the warm silver solu

Re: [Histonet] Bone marrow clot IHC tissue sections washing

[Regan Fulton via Histonet]

Martha,

We reported our study of 15 brands of adhesive slides for "wash off" and
found little difference among the different slides when well-fixed cell
culture material was examined.
On the other hand, poorly-fixed breast cancer tissues did appear to adhere
more strongly to some slides than others (TOMO being among the best).
Additional factors need to be considered, though, and I note that your
baking procedure is different from what is recommended by many slide
vendors.
In general, baking at 60-65 deg C for 1 hour is said to be optimal,
although we did not examine that parameter specifically.

Please see our poster at https://www.arrayscience.com/publications#Posters

Best regards,



Regan
Regan Fulton, M.D., Ph.D.
CEO and Co-Founder
Array Science, LLC
475 Gate 5 Road, #100
Sausalito, CA 94965
(415) 577-7360
email: ful...@arrayscience.com


www.arrayscience.com



On Thu, Sep 23, 2021 at 1:02 PM Martha Ward-Pathology via Histonet <
histonet@lists.utsouthwestern.edu> wrote:

> It was brought to my attention that we had significant washing on  3 of 8
> bone marrow clot sections the other day; this is not the first time so we
> would like to get to the bottom of this.   We use positively charged slides
> and all 8 cases were cut and run the same morning but allowed to air dry
> and then bake at 60C for 20 minutes before being run on our Bond 3
> stainer.   Has anyone out there experienced this type of problem and if so,
> what were your solutions?The repeat of the 3 cases today showed similar
> washing of tissue.
>
> This hasn't just started but has occurred periodically but the
> pathologists have tried to live with it and usually we can finally get
> enough tissue to stay on after 1-2 attempts.   Suggestions include cutting
> and drying the slides overnight and/or going to a gelatinated  slide versus
> a sialylated slide.   We have been using this particular brand of positive
> charged slide with good results for several years and rarely have issues
> with other tissue types unless they are particularly bloody.
>
> Thoughts or suggestions are greatly appreciated.
>
>
> Martha Ward MT(ASCP) QIHC
> Atrium Health Wake Forest Baptist
>
>
>
> ___
> Histonet mailing list
> Histonet@lists.utsouthwestern.edu
> http://lists.utsouthwestern.edu/mailman/listinfo/histonet
>
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[Histonet] Bone marrow clot IHC tissue sections washing

[Martha Ward-Pathology via Histonet]

It was brought to my attention that we had significant washing on  3 of 8 bone 
marrow clot sections the other day; this is not the first time so we would like 
to get to the bottom of this.   We use positively charged slides and all 8 
cases were cut and run the same morning but allowed to air dry and then bake at 
60C for 20 minutes before being run on our Bond 3 stainer.   Has anyone out 
there experienced this type of problem and if so, what were your solutions?
The repeat of the 3 cases today showed similar washing of tissue.

This hasn't just started but has occurred periodically but the pathologists 
have tried to live with it and usually we can finally get enough tissue to stay 
on after 1-2 attempts.   Suggestions include cutting and drying the slides 
overnight and/or going to a gelatinated  slide versus a sialylated slide.   We 
have been using this particular brand of positive charged slide with good 
results for several years and rarely have issues with other tissue types unless 
they are particularly bloody.

Thoughts or suggestions are greatly appreciated.


Martha Ward MT(ASCP) QIHC
Atrium Health Wake Forest Baptist



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[Histonet] Jones' Methenamine Silver Stain for Basement Membranes of Kidney - Issues and Questions

[Hood, Jordan via Histonet]

Hello,

I'm new to histology (and new to histonet), and I work in a small histology lab 
specializing in animal tissues that receives requests/submissions from 
researchers. I tried (and failed) to perform a Jones' Methenamine Silver stain 
on a client's submission of pig kidneys (formalin-fixed, paraffin-embedded, cut 
at 2.5 microns), and I need some help troubleshooting this stain since my 
co-workers are stumped, too.  I used the following procedure from Rowley 
Biochemical:


~
"Fixation: 10% Buffered Neutral Formalin (F-113) or Bouin's Solution (F-40) or 
Zenker's (F-155)

Sections: Paraffin, 2 microns

Procedure: Acid washed glassware must be used
1. Deparaffinize and hydrate to distilled water.
2. Oxidize in Periodic Acid 0.5% (F-396-1) 11 minutes. Wash in chloride-free 
water.
3. Prepare Methenamine Silver solution by mixing: 42.5 ml Methenamine 3% 
(F-396-2), 2.5 ml Silver Nitrate, 5% (F-396-3) and 12.0 ml Borate Buffer, pH 
8.2 (F-396-4).
4. Place slides in the solution and the entire jar in a water bath at 70°C for 
approx. 60-75 minutes. Check under microscope when slides appear medium brown 
microscopically. Every 10 minutes, once the medium brown color has been 
established, rinse a slide in 70°C, chloride free water and check under a 
microscope. Rinse again in hot water and return to the hot staining solution. 
As the staining time approaches the end point, check the slides, as above, 
every 1-2 minutes. The entire procedure must be performed quickly to prevent an 
uneven staining of the tissues. The slides should exhibit a brownish- yellow 
background, intense black reticulum fibers, and black basement membranes. If 
the slides become oversaturated, i.e. too black, destain in a dilute Potassium 
Ferricyanide Solution (F-396-11) for one or two dips.
5. Rinse well in distilled water. Tone in Gold Chloride 0.2% (F-396-5), 1 
minute. If sections are overtoned place in Sodium Metabisulfite, 3% (F-396-12) 
for 1-3 minutes. Rinse well in distilled water.
6. Sodium Thiosulfate 3% (F-396-9), 1-2 miutes. Wash in running tap water, 10 
minutes. Rinse well in distilled water.
7. Stain in Harris' Hematoxylin (F-396-6) containing 2-4ml of Glacial Acetic 
Acid per 100 ml for 5-15 minutes. Wash in water.
8. Differentiate in Acid Alcohol 1% (F-396-13) until the sections turn red.
9. Blue section in Ammonia Water, 0.3% (F-396-14). Wash thoroughly.
10. Counterstain in Eosin Y, 1%, Alcoholic Solution (F-396-7).
11. Dehydrate in 95% alcohol, absolute alcohol and clear in xylene 3 changes 
each. Mount.

Stain Results:
Basement membranes, reticulum fibers: Black
Nuclei: Blue
Cytoplasm, collagen, connective tissue: Pink-orange

References: Jones, D.B., Amer.J.Path. 27:99 (1951). AFIP Manual of Histolocical 
Staining Methods, 3rd ed., Ed. L. Luna: NY: McGraw-Hill Publications, c. 1968, 
p. 97."
~


It became apparent that something went wrong during Step 4 when the slides were 
in the glass container (not a coplin jar - we have ten slides that we need to 
stain so we're using a rectangular glass container that holds ten slides on 
their sides - it does require a metal handle to move, but the handle is 
flexible and easy to remove after the glass slide rack has been transferred 
between containers) of silver solution in the water bath because there was lots 
of precipitate on the slides and floating on the surface of the silver solution.

In my first test, I used five test slides (extra slides that we cut from the 
same blocks that were submitted to us). I deparaffinized them in coplin jars 
(moving them with plastic forceps) and hydrated them to deionized water. I 
transferred the slides to a glass slide rack that holds ten slides on their 
sides, added five blank slides that were rinsed in deionized water (so that the 
displacement of reagents would be equivalent to when we stain our ten "real" 
slides after testing is complete), and completed Step 2. I don't recall exactly 
how long the glass container of silver solution and the glass container of 
deionized water had been heating up in the water bath, but I would estimate 
~15-30 minutes. The thermometer said that the water in the bath (not inside the 
containers) reached ~60-65 degrees Celsius. The silver solution was clear and 
colorless when I made it up, but by the time I put the slides into the warm 
silver solution, the solution was beginning to turn a light brown color (though 
it was still clear and I did not see any precipitate floating around). I 
removed the metal handle of the glass slide rack after the rack was transferred 
into the silver solution, but the metal handle did dip into the silver solution 
briefly. At some point, I noticed precipitate floating around of the surface of 
the silver solution. After ~80 minutes, I used plastic forceps to remove one 
test slide from the warm silver solution, dipped it several times into the warm 
deionized water to rinse it, and wiped off the back of the slide with gauze. 
The amount of precip

Re: [Histonet] Multiple H&E stainer maintenance

[Bacon, Charles via Histonet]

Hi Leslie,

I have attached our current log. You can augment it based off the solutions you 
are using and how many shifts you have. We have two stainers and each has their 
own log. Our stainers are connected to running water thus the drain lines that 
need cleaning. If you have any questions reach out anytime. Good luck!

Best,

Chuck Bacon, HTL(ASCP)CM 
Supervisor Histology
Baystate Medical Center
361 Whitney Ave., Holyoke, MA 01040
Telephone: 413-322-4786  Fax: 413-322-4790
charles.ba...@baystatehealth.org

-Original Message-
From: Hanson, Leslie I :LLS Lab [mailto:lihan...@lhs.org] 
Sent: Wednesday, September 22, 2021 2:29 PM
To: 'histonet@lists.utsouthwestern.edu'
Subject: [Histonet] Multiple H&E stainer maintenance

Hi all,

We recently acquired a second H&E stainer and are working up a maintenance log. 
Looking for input from others who have already tackled this issue!

  *   Do you have a log for EACH instrument or one combined log?
  *   Do you have a space for listing reagent lots?
  *   What is your rotation schedule for changing reagents?
  *   Other tips and tricks are always appreciated!

Thanks!


Leslie Hanson, HT(ASCP)
Tech Specialist
IHC / Pathology
Phone: (503)944-7923
lihan...@lhs.org



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