RSS Feed Print
BOD/CBOD toxicity??
runnerlabber
Posted: Tuesday, February 9, 2010 1:51 PM
Joined: 1/26/2010
Posts: 6


We often will have BOD/CBOD results that make us wonder if we are reporting the result correctly. So I was wondering how others might report the BOD results based on the data as follows (all samples were seeded):

 

#1. Primary influent  sample vol.   init DO   final DO   BOD

     (CBOD)             1 mL            8.49       6.49      280

                            3 mL            8.39       5.84      148 

                            5 mL            8.22       4.83      140 

 

#2. Primary influent  1 mL            8.43       5.89      410

     (BOD)               3 mL            8.30       4.72      240

                            5 mL            8.12       2.66      264

 

#3. Primary influent   1 mL            8.73       6.21      252

     (CBOD)              3 mL            8.60       5.33      159

                            5 mL             8.38       4.38      162

 

                         


Keith Chapman
Posted: Tuesday, February 9, 2010 2:49 PM
Joined: 2/5/2010
Posts: 5


before citing toxicity consider how you pipetted the 1 ml volume.  I doubt you have real toxicity as the two larger volumes both have quite similar BOD's.

 

I suggest adding the 1 ml volume by first making a 1:10 dilution by adding 10 ml of sample to a graduate and bringing it up to 100 ml of volume with DI water, then pipet 10 ml of this dilution into the BOD bottle.

 

This might solve the apparent toxicity issue.  Perhaps.


runnerlabber
Posted: Tuesday, February 9, 2010 3:07 PM
Joined: 1/26/2010
Posts: 6


Actually, I did not include that this sample is prepared as a 1:10 dilution to help make it more homogeneous. So we put in 10, 30 and 50 mLs into each bottle.


James Royer
Posted: Tuesday, February 9, 2010 3:23 PM
Joined: 9/21/2009
Posts: 98


What is the seed correction as I can not figure how the BOD results were calculated?


Perry Brake
Posted: Tuesday, February 9, 2010 4:05 PM
Joined: 12/16/2009
Posts: 69


According to the method, you should average the three results for each sample.  Unfortunately, many permit managers and even lab certification/accreditation assessors wouldn't be interested in hearing something like, "the 1 mL bottles...which are really 10 mL...just barely depleted 2.0 mg/L of DO, so I didn't include them in the average" which is the most logical thing to do with the results you posted, in my opinion.  All they would say is, "You need to record what you actually did [i.e., analyzed samples of 10, 30, and 50 mLs of a 1:10 diluted sample] and average all valid results."

 

What I would recommend you do is forget about the 1.0 mL (or 10 mL of the 1/10 dilution) and do 3, 5, and 10 mL of an undiluted sample.  You might find that the 10 mL dilution sometimes fails to leave 1.0 mg/L DO, but in those cases, the method just tells you not to include it in the calculations.  You would be legal, and your results would be just as valid, if not more so.  To get a homogeneous aliquot, rather than diluting, you can blend it (at low speed...high speed might injure natural seed bacteria) or vigorously stir it, and take the aliquot from the sample immediately after turning off the blender/stirrer using a wide-mouth pipet.


runnerlabber
Posted: Tuesday, February 9, 2010 4:15 PM
Joined: 1/26/2010
Posts: 6


Example  #1 had a seed factor of 1.07 mg/L

#2 SF was 1.18 mg/L

#3 SF was 1.68 mg/L

 

Hope this helps.


runnerlabber
Posted: Tuesday, February 9, 2010 4:21 PM
Joined: 1/26/2010
Posts: 6


Thanks Perry. We actually do show on the benchsheets that the sample is diluted 1:10 so as not to be hiding anything.


James Royer
Posted: Thursday, February 11, 2010 7:53 AM
Joined: 9/21/2009
Posts: 98


The first thing I would do is check the COD of my sample to determine the estimated BOD. If the COD is 300 then maybe the 150 BOD is correct and the 400 BOD would be an obvious bad result.

My first thought is that the seeding material might contain nitrifiers and the inhibitor is being overwelmed. The highly diluted samples would then supply more ammonia to be oxidized in relationship to the organics in the sample.

I would try using less seed material lowering the seed correction factor, analyzing a G-GA standard, and trying the analysis without seeding the primary influent. After trying these you can reevaluate the problem. Jim


Anonymous
Posted: Friday, February 19, 2010 3:50 PM

So how would you report the following results from an unknown source:

 

5 ml of sample (diluted 1:10) = 729 mg/L (DO depletion 2.6)

10 ml of sample (diluted 1:10) = 535 (DO depletion 3.17)

30 ml of sample (diluted 1:10) = 574 (DO depletion 7.12)

 

if I read all comments correctly, including "The Bug's-Eye-View of the BOD Test", I could:

a. regard the first result as an outlier, mark it as such and report the average of the last 2 results

b. report the average of all three results

c. consider this as possible toxicity since the difference between high and low results is greater than 30% (Standard methods), report the highest results and mark it as "possible toxicity".

 

Any suggestions?


Perry Brake
Posted: Saturday, February 20, 2010 12:35 AM
Joined: 12/16/2009
Posts: 69


 

I don't think you found all of the commentary in choice "c" in A Bug's-Eye-View...you don't have enough results to report the sample as showing toxicity (the author of A Bug's-Eye-View suggests doing at least seven bottles). 

 

You can't go wrong with choice "b" because that's what the method says to do.  On the other hand, if you think you could convince your regulatory agency that the first bottle is unreliable, if for no other reason than it barely depleted the minimum 2.0 mg/L, you might want to report "a."  I don't think any regulator would ever find fault with that...unless they really don't understand the BOD test and insist on you following the letter of the law.  And even they would probably just say "don't do that again" and that would be it.

 

If you really suspect toxicity is having an affect, add four more bottles to your run, diluted 1:100 and doing maybe 30, 20, 10, and 5 mL dilutions.  If the BOD values continue to increase as the bottles become more dilute, you could defend claiming toxicity and reporting the highest result for the sample that depleted at least 2.0 mg/L while leaving at least 1.0 mg/L.


James Royer
Posted: Monday, February 22, 2010 10:43 AM
Joined: 9/21/2009
Posts: 98


I would still check the COD to know what the maximum BOD should be. I would not be comformable throwing out you 700 BOD on the 5 ml when the 10 ml result was higher than the 30 ml result. Perry is correct to indicate that you need to have more dilutions to further evaluate what is happening and justify discarding erronous results.


James Royer
Posted: Monday, February 22, 2010 10:45 AM
Joined: 9/21/2009
Posts: 98


OOPS. 30 ml BOD results higher that the 10 ml BOD results were what I meant.


Anonymous
Posted: Saturday, March 6, 2010 7:38 PM

I am new to BOD analysis in an environmental lab. Recently encountered an example with a COD of ~1500 mg/L (repeated on 3 occasions - each time confirmed) and a BOD using 3 dilutions with very consistent results of close to 4000 mg/L.

By the time this discrepancy was discovered the sample was out of BOD holding time by nearly a week. An attempt to nevertheless repeat the BOD using similar dilutions failed to generate the same, and also not at all consistent results, so figuring out what was possibly going on here is very difficult.

Will appreciate it if anyone can offer a possible explanation/s.

I will have to look on Monday if the sample contained high NH3 - could this be a possible cause of such a big discrepancy? If not, what else could have caused BOD >>>> COD?


Perry Brake
Posted: Saturday, March 6, 2010 10:37 PM
Joined: 12/16/2009
Posts: 69


You say you used 3 dilutions to come up with the 4000 mg/L BOD.  It would help if we knew what those dilutions were, and what depletion and seed correction was associated with each dilution.


Anonymous
Posted: Monday, March 8, 2010 1:40 AM

Hello Perry,

 

Due to a very strong smell (sulphury?) it was given an initial dilution of x50, then 3 secondary dilutions of 10, 20 and 50 mL per 400 mL.

Overall initial plus secondary dilutions as follows:

0.05% dilution depleted form 8.32 to 5.82, calc'ed as 3884 mg/L

0.10% dilution depleted from 8.30 to 3.82, calc'ed as 4120   "

0.25% dilution depleted from 8.25 to 0.26, cal'c as 3203    "

 

The 3rd dilution depleted too much, so would be disregarded, but still gave an indication of the BOD being high.

 

At the same time the COD was only ~1500 mg/L.

 

Little else was analysed, so I don't have other data to draw on.

Using simple test strips I can see the pH is neutral, it contains no nitrates, but it does give a positive reaction for ammonia. (around 6ppm or higher)

 

I'm struggling to make sense of this.

 

Can the presence of ammonia elevate the BOD to this extent?


Perry Brake
Posted: Monday, March 8, 2010 1:04 PM
Joined: 12/16/2009
Posts: 69


 You saw through my line of questioning about the dilutions...without a dilution of the whole sample, it would be rather difficult to come up with BODs in the thousands.  Good move on your part!

 

Presence of ammonia will result in higher BOD results while not affecting the COD results, but not to the extent you are seeing. If you still have some sample left...or have a very similar, sulfur-smelling sample available...you could do both a BOD and CBOD and get a feel for the contribution of the ammonia or other reduced forms of nitrogen.

 

You didn't mention the blank(s) for that batch.  Because your total dilution factor is so high (e.g., 1500 for the 10 mL dilution...I am assuming that "400 mL" is supposed to be "300 mL"?) if the source or dilution water (as opposed to a contaminated blank bottle) causes a high blank, it carries over into every bottle that uses dilution water.  For example, a 0.1 mg/L blank could increase the BOD result by 150 for the 10 mL dilution.  When you made your initial 1:50 dilution of the entire sample, were you careful not to supersaturate the solution?  If supersaturated, it is conceivable that the water could be causing very high errors, again because of that huge total dilution factor.

 

Sulfides (which are causing the smell) have an oxygen demand and are undoubtedly causing part of the high BOD results, but they are also oxidized in the COD test, so they probably are not a major contributor to

 

 

 

the disparity you are seeing between BOD and COD results.

 

I hope somebody else will speak up on this anomaly...it sure has me wondering!


James Royer
Posted: Tuesday, March 9, 2010 8:53 AM
Joined: 9/21/2009
Posts: 98


There is no indication of toxicity but values need to be verified before any data should be reported. The COD of about 1500 needs to be rechecked as it might be at the limit if analyzed by the Hach method. A dilution should be run on the sample to verify that the COD is correct.

I always preserve a portion of unknown samples with H2SO4 so that the COD's can be rechecked. That same sample can then be neutralized and seeded so that BOD can be rechecked also.

If the sample looked that bad it would be wise to seed the BOD analysis to ensure that there are enough bacteria to utilize the organics in the sample. This does not seem to be the problem if the BOD's are higher than COD. A seeded sample also allows for the standard G-GA sample to be analyzed so as to verify that the dilution water and analysis is good. Setting up both T and C BOD will allow for evaluation of ammonia interference.

If the sulfide is very high and you preserved the sample for COD the sulfides would have evolved off and not shown up in the COD value. Make a dilution and run both BOD and COD from the same dilution. A history of the sample would be helpful.


Luis Manriquez
Posted: Tuesday, March 16, 2010 6:17 PM
Joined: 1/6/2010
Posts: 20


Our BOD software has built in rules for handling replicates like these and we also manually apply additional rules.

 

The rules are:

A- If there are 3 results and they are within 20% of each other, we average all 3.

 

B- If there are only 2 results within 20% we average those 2 and ignore the outlier.

 

C- If there are only 2 results and they are over 20% apart, we use the result with the largest DO depletion and qualify the data.

 

 

 

D- If there are 3 results and they are over 20% from each other and decline with increasing concentration, we use the result with the smallest sample volume (highest BOD) and qualify the data.

 

 

 

E-  If there are 3 results and they are over 20% from each other and do not decline with increasing concentration: Check bottle sequence. Check COD results and discard any whose BOD are more than the COD.

 

F- If the BOD result is less than 10, do not use the 20% rules. Average all valid BODs.

 

In the examples given, the rule B appears to apply. We have a nice decision tree made up in Excel that shows all the possible outcomes and aids in making the correct decision.

 

 

 

 

 


James Royer
Posted: Wednesday, March 17, 2010 8:18 AM
Joined: 9/21/2009
Posts: 98


Luis has covered the BOD reporting in a very good manor. He has a writen procedure to follow so that all analysts, and inspectors, can follow what and why data was used or discarded. If there is a change in the standard procedure they can simply update their reporting proticall. Inspectors should recommend this and maybe Standard Methods can include this in the method.


Luis Manriquez
Posted: Thursday, March 18, 2010 4:54 PM
Joined: 1/6/2010
Posts: 20


James:

Thanks for the thumbs up.

 

 

I should have added that we always divide the smaller result by the larger result on each comparison pair when determining if within the +/-20% limit.

 

Also, the qualifying statements we use are:

 

C.- "Result from largest DO depletion reported."

 

D.- "Possible toxicity. Highest BOD reported."

 

E.- "Result from largest DO depletion reported."

Note that E is a special case and if the cause is not found then applying the rule from C is suggested.

 

These rules were in my software from Man-Tech, but they are pol\pularly called Texas Rules. Does anyone know how these originated?

 

Luis Manriquez


Luis Manriquez
Posted: Friday, March 19, 2010 11:08 AM
Joined: 1/6/2010
Posts: 20


Perry:

Greetings from (mostly) sunny Arizona. I could not reply to your e-mail since it came from the web administrator. but I thought you would not mind if I replied here.

Your comment that C and E are identical is not quite correct. Rule C applies when there are only 2 results and they are over 20% apart.

Rule E applies when there are 3 valid results over 20% apart and they do not decline with increasing sample volume.

If you find a sample with the E situation, you need to investigate what happened as it is an unusual result.

 

While applying the rule from C to an E situation is one option available, it is not mandatory.

Perhaps one may find that the bottle sequence was switched or that one or more bottles were higher than the COD result, indicating a bad sample set up.

Other possible causes might be transcription errors, i.e. an incorrect volume entered in the spreadsheet than what is in the benchsheet.

And there are still other options.

For example, one might decide to use the result closest to the historical data, if available, with a qualifying comment.

 

Thanks for your feedback. I really enjoy this.

Luis

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 


Perry Brake
Posted: Friday, March 19, 2010 1:21 PM
Joined: 12/16/2009
Posts: 69


I don't know what I said, but what I MEANT to say is the two qualifying statements, C and E, are identical.  But  you...and probably everyone else reading it...knew that!

 

 

You're right...it is fun...but not nearly as much as it was when thousands were posting on this technical forum.


Anonymous
Posted: Tuesday, April 20, 2010 11:07 AM

If you subtract you seed amounts from your sample depletions for the 1 mL dilutions, your residual DO is less than 2 mg/L.  I would not use any of these dilutions for the calculation of the sample result.  Standard Methods says that you must have sample depletion of at least 2.0 mg/L.  We have always interpreted this to mean that the depletion must be from the sample and not the seed.  These results are at the very low end of the ability of this method to detect concentrations reliably.


Perry Brake
Posted: Tuesday, April 20, 2010 4:26 PM
Joined: 12/16/2009
Posts: 69


The 21st Ed. of Standard Methods changes that and says that the minimum 2.0 mg/L depletion includes the seed.


James Royer
Posted: Thursday, April 22, 2010 8:14 AM
Joined: 9/21/2009
Posts: 98


Perry is correct, as both the 20th and 21st ed. of Standard Methods are clear that "for each bottle" the depletion must be 2.0 mg/L depletion. This is why it is important to try and keep the seed correction value to .6 to 1.0 mg/L as greater than 1.0 would be over half of the depletion if only 2.0 depletion occured.


Luis Manriquez
Posted: Thursday, April 29, 2010 10:45 AM
Joined: 1/6/2010
Posts: 20


Mr. Royer:

 

You have answered a question I had for a long time: How did SM arrive at the 0.6 -1.0 mg/l limit for seed depletion?

Now I know the rationale for these numbers. Thanks!!

 

Luis Manriquez