Science topic

Bronchoalveolar Lavage - Science topic

Bronchoalveolar Lavage is a washing out of the lungs with saline or mucolytic agents for diagnostic or therapeutic purposes. It is very useful in the diagnosis of diffuse pulmonary infiltrates in immunosuppressed patients.
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We are planning to study inflammatory cells in mouse bronchoalveolar lavage fluid. Briefly, we would perform the bronchoalveolar lavage, collect the fluid, centrifuge it, freeze the supernatant at -80 degrees C with proteinase inhibitors. Then we would lyse erythrocytes with ACK lysis buffer for 2 min., dilute the ACK buffer with PBS, centrifuge again.
I have two questions:
1. Should we add EDTA to prevent cell clumping?
2. How should we perform the Wright-Giemza stain? We do not have a cytocentrifuge. Can we just resuspend the cell pellet in a small ammount of PBS and add the cell suspension onto a slide to dry out and stain? Maybe someone has a similar protocol that you could share?
Thank you!
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You need to have some method of centrifugation for the cells to stick to a slide and get a bit flattened out so that their features can be evaluated microscopically. If you don't have a cytospin, there is another technique which involves a basic centrifuge. Use cylindrical flat-bottom tubes, inside which you place a round slide coverslip of just under the diameter of the tube. Then put the BAL fluid in the tubes and centrifuge. The cells will stick to the coverslip, which you then remove from the tube and stain with your method of choice, and then mount the coverslip cell-side down on a standard microscope slide.
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I need to isolate alveolar macrophages from ~20 ml bronchoalveolar lavage (BAL) of patients with pulmonary tuberculosis. The BAL was filtered and then centrifuged for 10 min, 250g, 4oC and wash 1x with RPMI. Then I put the cells into 48-well plate in RPMI + 1% L-glutamine +10% fetal bovine serum +1% pens/strep + 1% puruvate and incubated at 37oC, 5% CO2 o/n. About 20% cells were died when I checked by tryphan blue after isolation. The day late, I washed out to remove the non-adherent cells. Most of the cells were washed out, very rare cells could adhere to the surface. Does anyone get the same problem with me? What should I need to change to improve the viability of cells as well as percentage of adhered cells? Thank you!!
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Hey, I hope that someone can help me, I want to extract cells from BALF but we dont won a cytospin in our lab, could we do it otherwise ?
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bronchoalveolar lavage sample
the symptoms are fever shortness of breath and tender lymph nodes
I thought it may be toxoplasmosis but this image is at x100 magnification and the pictures dont match
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Toxoplasma gondii tachyzoite
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I am looking to purchase an Automated Cell Counter that can also do asses viability/cell differential for our lab. We do lots of cell counts/differential for bronchoalveolar lavage cells, blood cells and cell culture applications.
Looking for recommendations for a solid device with good customer service/trouble shooting resources.
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You can barrow three part or five part cell counter for you parameters
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After performing bronchoalveolar lavage in mice lung , i don't see any inflammatory cells in my sample , do you have any Idea ? Maybe the problem is in my PBS ? Can i use normal saline instead if PBS? Or the problem is in the coloration ,thanks
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Reshed Abohalaka thank you sir, but how can have my cell pellets without using a centrifuge?
do you have any idea if I can use normal saline instead of PBS ?
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670 / 5000
We are conducting a standardization of our fungi DNA extraction and have made serial dilutions of fungi conidia in bronchoalveolar lavage (BAL - negative for fungi and bacteria).
We added conidia to the BAL and calculated concentrations of 1,000,000, 100,000, 10,000 conidia / mL.
However, on several occasions, our bands with different conidia concentrations appear in different sizes, with the sample containing the highest concentration of conidia (10 ^ 5) being lower and the next ones being higher, successively. (10 ^ 4 below 10 ^ 3, 10 ^ 3 below 10 ^ 2).
Has anyone seen this happen and what is the possible reason?
Edit: The bands are from PCR product with fungi primers (it happened with universal fungi primers and also with specific primers).
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It may be that in the dilutions and then addition of sample to the pcr mix you are changing the salt concentration of the sample and then the pcr mix.This Might change the running speed of samples so that the highest salt concentration causes the slowest moving band through the gel
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Hello. The question I wanted to pose to you is concerning in vivo assays. One of the possibilities of assays is a bronchoalveolar lavage (BAL). This technique entails que lavage with PBS (for example) of the airways, and collect that fluid, and further analyse it (proteins, enzymes and cells).
In your expertise, can we store the liquid from BAL? Like freezing, for example.
Or do we have to analyse right away?
Do you have any advice on how to tackle this?
Thank you all for your help in advance!
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Hi Jorge,
We have 2 policy for this issue. If BAL analysis and cytology is about to done in hours, we send fresh fluid (BAL) to Lab. For those specimen, which maybe evaluated after longer period, we routinely add equal amount of 96% alcohol (50, 50) as cell fixator.
Thanks
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Hello dear community,
currently I am facing some issues with a bacterial murine model of pneumonia with Klebsiella. To access the signs of lung injury is straightforward with counts of lung lavage CFUs. However, further evaluation of BAL by flow cytometry (i.e. phagocytosis, transmigration, adhesion), ELISA and other techniques exposes the lab. staff and equipment to persistent contamination with a high risk of development of human pneumonia.
Murine models are very well described in the literature but human health low-risk alternatives are not. For our future work, we are considering between the most common sources of murine pneumonia, bacterial or viral, and trying to understand which one has less human health impact in the lab. For the bacterial alternative I have considered Bartonell henselae, Klebsiella pneumonia, Pasteurella pneumotropica, with the first being the safest. Conserning the Pneumonia virus in mice, the family Paramyxoviridae, subfamily Pneumovirinae seems to be the standard but the literature is limitless.
Your advice will be extremely valuable for our future experiments and safety of our lab. members.
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Thank you so much for your answer.
The effort I am trying to make is to avoit complete exposure of lab. staff to contaminants. A S2 lab. would be enough to contain any bacterial or viral infection. But since we are integrated into a medical school, attendees are not aware of the risk of working with pneumonia and spread easily contaminants.
To work with a virus that would infect only rodents would be ideal. Working with a bacterial species that would only infect immuno-compromised can be handled.
All the best.
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Hi
Does anyone have a detailed protocol for bronchoalveolar lavage (BALF) in rats? So far my LIVE/DEAD cell ratio in the BALF is very low (20%/80%).
Also can anyone share a Wright-Giemsa staining protocol (Cell differentials) in the BALF?
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the attached docs give a detailed method
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I've tried to lavage the lung via trachea with phosphate buffered saline (PBS) however after I inserted the PBS into the lung it was difficult to retrieved it back and sometimes the fluid inserted was leaking.
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@ James Leigh thankyou for your response
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Hello everyone!
I would kindly ask if there is someone who has some experience in taking BALF from mice. I´ve seen people taking BALF from deeply anesthetized mice but my impression is that most of the people are taking the BALF from dead/exsanguinated mice.
Are there any differences between these to ways regarding the cell-count or protein-level in the BALF? What would you recommend?
Many thanks in Advance!
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Anesthetized is better. You can use the following protocol:
- Prior to each allergen challenge, and following the 7th challenge, mice are administered an intraperitoneal injection of 48 mg/kg etomidate (2 mg/ml), prior to placement in a light-excluding receptacle.
-Subject remains in receptacle until a lack of observable neurological response is detected upon application of pressure to hind paws (5-10 min).
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1. Why must cytospin smears be air-dried and unfixed?
2. Won't the cells morphology and contains change if the cells gets dried? Can they still be differentiated after staining?
3. Actually we don't have a cytospin in our lab and I want to spread the cells on the slides without using Cytospin. So could I fix the cells first and then get them air-dried?
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According to my experience, it's quite difficult to prepare slides from BAL fluid without cytospin. Some crust-like formation happens when the slide is left for drying on its own due to evaporation of the buffer and cells don't stick that well onto the slide. That's why cytospin is used with a filter paper to absorb extra fluid and impact only the cells to the slide. Overlapping of  cells is also a problem as you need to put a concentrated sample onto the slide. Therefore differential cell count becomes a problem with such a method.
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Can anyone advise me on an efficient tool to make broncho alveolar lavage on rats? I usually do it on mice with a catheter, but I would like to do the same on rats and I don't know which tool pick and which size?
Thanks a lot for your answers
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Within 2 weeks, I will perform bronchoalveolar lavage cytospin on Superfrost Plus slides, and then make a Diff quik staining.
There is no Diff Quik staining available in lab for the moment and I can't receive them before 1 month. Does anyone know how to stock the slides meanwhile the kit arrive?
Thanks in advance!
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The sooner you stain the slides the better but if that is not an option try at least to fix them ASAP. The first step of Diff Quick is Methanol (that is the fixation step) so you can purchase methanol (or maybe you have it in your lab) and fix the slides.. once you get the Diff Quick can proceed with steps 2 and 3... Also Merck sells a Diff Quick alternative (Hemacolor) that works very well.. maybe you can check with them if they have it..
Good luck on your experiments
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I was using a metallic probe to have access to the trachea and I inject 500µl of PBS (Phosphate-buffered saline). 
Using the Biorad Milliplex Kit to quantify the cytokines in BAL, I have almost nothing in BAL that's why I'm wondering if there was a problem when I perform my BAL.
Do you have any recommendation to optimize my protocol?
Please help! Thank you..
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See the article attached "Simvastatin Pretreatment prevents ambient particle-induced lung injury in mice". The standard protocol used in the lab and described in the article stands for 12 lavages of 1 ml each with cold PBS with a metallic probe. However if you are quantifying cytokines in BAL, you only need to dose the first ml lavaged. The result also depends on which cytokine are you studying and the time passed after the stimuli/insult in your experimental design, as every cytokine has an expression profile, sort of a temporal window where they can be detected. Maybe in your experimental design, you have to dose serum instead of BAL to check the systemic levels.
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I am using the Biorad Bio-Plex Pro assay to quantify different cytokines in mice BAL (Bronchoalveolar Lavage), plasma, and lung samples.
I have no idea if I should add proteases inhibitors to avoid proteolysis.
Do you have any advice or idea to optimize the quality of the samples?
PS: I have already BAL and lung samples stocked at -80°C without proteases inhibitors, is it a good idea to add them just before the multiplex experiment? (in case proteases inhibitors are obligatory)
Thanks in advance!
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Hi Yara, it is always a good idea to add proteinase inhibitor cocktail to biological samples kept at room temp.
In general, all proteinases are inactive at -80oC. (they are hydrolases that require free H2O and at -80oC, there is almost no free H2O). 
Some are active at -20oC, but all are active at +4oC to +25oC.
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The featured layouts are from 3 adult BALB/c mice that have been infected with RSV Line 19 for 7 days. The BAL was stained for CD11c and CD11b for flow cytometry. The CD11c- CD11b- granulocytes (appear as blue on FSC vs. SSC) make up as much as 60% of the granulocyte gate. The naive group's granulocyte gate from the BAL appropriately stained as 97% CD11c+ CD11b-, so we know that it wasn't a staining issue. Any insights are appreciated!
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Check it these cells are CD45+, if positive - stain for Siglec F (eosinophils), if negative - check for EpCAM, if positive - probably AT2. Also, no such thing as a granulocyte gate: it is 21st century, we should be defining cell populations via markers, not via FSC/SSC. 
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I would like to generate human PBMC from BAL and to stimulate them (specially Treg) with PMA ionomycin on 3-4 hours.
Thank you for your help !
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Hi,
PBMC stands for peripheral blood mononuclear cells, so by definition you cannot isolate them from BAL, unless there is blood in the lavage.
Could you try to be more precise? Is it monocytes you want? Or leukocytes? Or lymhocytes?
Kind regards,
Agnieszka
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About bronchoalveolar lavage in mice.
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identify the trachea then make a slight horizontal incision, make sure you dont disconnect the trachea. insert a 16 gauge needle and ligate the whole unit. inject 2 ml of freshly prepared PBS and collect it back and store it at -20 oC till further estimations. Make sure you use 2ml and not more than that
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Observing a very frothy white layer on a bronchoalveolar fluid obtained with a very  good yield (155/200ml) prompted me to measure both the albumin and protein content of this fluid. The beer-like appearance reminded me of dissolving human albumin some years backi in a lab experiment. The concentrations obtained were very low (measured with the technique used for detection of microproteinuria...).
Protein 0.132 g/L and albumin 11.2mg/L. So far I have not found reference values for these findings.
Anyone aware of the normal range of protein and albumin content in BAL fluid?
Is the froth related to the yield rather than to an increased protein content?
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READ ABOUT THE UREA METHOD...
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I'm looking for a protocol for MGG staining protocol for BAL and peritoneal cells smears for the morphological assessment of the different cell populations.
Is it the same as the standard protocol used for blood smears?
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Get your BAL and peritoneal washings unfixed. Make cytospins and air dry the spins. Then try an ordinary MGG protocol. You can make adjustments to the staining in the washing procedure at the end, if necessary. You should filter your Giemsa solution often, to avoid staining particles to pollute and disturbe the cellular materiale.
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Last week a new HIV patient was admitted to the Infectious Diseases Unit where I'm currently working; he's coming from Pneumology where he was admitted for respiratory insufficiency. PCR from his sputum was negative for P. jiroveci, while it was positive for CMV-DNA (3 x 105); his CD4 were 5, HIVRNA 90170 cp. Clinical findings were suggestive for PCP and when he was transferred to our Unit we started treatment for PCP (TMP-SMX IV, corticosteroids, O2) and ARV therapy; the day after starting ARV we performed a BAL : PCR positive for CMV-DNA (2,5 x 108), positive forEBV-DNA (107), positive HHV6.
Some colleagues of mine think we have to start antiCMV therapy (Gancyclovir), others suggest to attend ( they say that CMVDNA positive on BAL is not diagnostic for a CMV infection).
What do you think about?
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Dear Goffredo,
Treat the PCP
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I have seen 2 cases of PAP in dogs. Both were near endstage disease and lung biopsy not possible. BAL samples were considered normal. PAS and Papanicolaou stains are not typically done in veterinary medicine. Are these the two stains most beneficial for diagnosis of PAP from BAL samples. Is there a naturally occuring animal model?
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Normally the BAL should look milky white due to the masses of protein debris. We do only May-Grünwald-Giemsa staining. In this case you will see foamy macrophages on a "dirty" background. This dirty background consists of acellular granules of protein debris that may stain intensely blue. Sometimes these granules are rather large. These acellular granules are also PAS positive. We never did Papanicolaou staining.
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I am trying to collect Bronchoalveolar lavage from mice. I learned it from another lab where they used needles to insert into the trachea. But when I was trying, the liquid could leak and the needles sometimes got out of the trachea. I checked the protocols online and I found lavage tubes were used in some protocols.
Could anyone who is using lavage tubes to collect BAL from mice, please provide me wit further details (supplier etc.)?
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We use a 22ga x 3/4" Luer adapter as a tracheal cannula for our BALs. It's essentially a 22ga blunt stainless steel needle with a female Luer hub that allows you to attach a syringe for injecting the lavage fluid. Here is our BAL protocol for mice:
Make a 1.5-2cm longitudinal incision on the ventral side of the neck.
Expose the trachea by blunt dissection a work curved forceps underneath the trachea.
Use the forceps to pull a 10-12cm piece of suture thread underneath the trachea.
Pull on the ends of the suture, cranially, to "stretch" the trachea in front of you.
Use a 20ga needle to poke a hole in the trachea as close to the larynx as possible.
Insert the 22ga cannula into the hole and advance approximately 5-8mm (stop short of the carina to avoid puncturing or tearing the bronchi).
Secure the cannula in the trachea by tying the suture around it with an overhand knot (the reason we use the steel cannula is so we can tighten the suture knot snugly to prevent leakage).
Attach a 4-way stopcock to the cannula with 2, 5cc syringes attached to it (one contains 5ml of lavage fluid and the other is empty, ready to collect the recovered BAL fluid).
Alternate injecting 1ml of fresh fluid and collecting 5X. Obviously, you can adjust the volumes based on your particular needs.