Category Archives: Antimicrobial Resistance

Plasmids and Team Players

Let’s say you have a problem at your hospital with carbapenemases.

One of the obvious solutions would be to reduce the use of carbapenems in order to reduce the selection pressure.

However even if you stopped carbapenem usage altogether the carbapenemases would not necessarily disappear…

This is because carbapenemases are often plasmid borne, and there are often antibiotic resistance genes for other antibiotics, e.g. A, B & C sitting on the same plasmid.

As long as the (high) usage of antibiotics A, B & C continued then the selection pressure would favour plasmid retention in the bacterium, and thus allowing persistence of the carbapenemase.

Selection pressure by proxy.

Are we all doomed?

Not necessarily…

A gene expressing one antimicrobial resistance determinant comes at an energy cost to a bacterial cell. Plasmids expressing multiple resistance genes come at even more energy cost to the cell. You can be sure if it did not need the plasmid to ensure its survival, it would be mercilessly dumped, and probably sooner rather than later.

Therefore even a modest reduction in carbapenem usage, along with a reduction in antibiotics A, B & C may go a long way to solving your problem.

Advances in molecular methods and whole genome sequencing over the next decade will mean that it will become much easier to work out exactly which resistance genes are contained in the plasmids circulating in our local hospitals, and anti-microbial stewardship can thus be optimised accordingly.

Sounds space age?

Not really, we just need to be aware that resistant bacteria are very smart in an evolutionary sense, and we need to stay alert, and not give them the niches they are looking for…

Michael

Illustration courtesy of www.biologyfun.blogspot.co.nz

“Keeping it simple or keeping it accurate…”

Let’s say you are a clinician and you are looking at two different microbiology results on two different patients (A&B). Both have an E. coli UTI. Both results state that the E. coli is susceptible to trimethoprim. However what you don’t know is that the E. coli isolate on Patient A had a trimethoprim disc diffusion zone of 18mm (right on the EUCAST breakpoint), whilst for patient B the corresponding zone was a much more comfortable 26 mm.

And who knows, if you repeated the same testing on patient A a dozen times, the chances are you would have a few “non-susceptible”results, due to the natural margin of error of the test.

If I was a clinician, and had this extra (zone diameter) information, I would be a lot happier prescribing trimethoprim to patient B, even though they both have in-vitro “susceptibility” reported on the result. (The same principle of course applies if we were talking about Minimum Inhibitory Concentration (MIC) values instead of zone diameters.)

But do clinicians really want this extra information?

They are usually very busy, …and not particularly interested in microbiology.

In my experience all clinicians generally want to know is if an isolate is susceptible or resistant. They are not particularly interested in the details, with the exception of blood culture and sterile site isolates, when there is at least a modicum of interest in the degree of susceptibility or resistance.

So which is better.. a susceptibility result full of information, but potentially difficult to understand and interpret, or a result reduced to its simplest form.

There is no right answer of course…

I am not even convinced antimicrobial susceptibility breakpoints have a long term future.

More and more, year by year, the anti-microbial susceptibility committees (EUCAST, CLSI) are trying to take into account antibiotic dose,  renal function, degree of infection, etc. when setting antimicrobial breakpoints.

But they are really only scratching the surface…

Time for some major disruption!

Michael

“Trending…”

I get the occasional anxious phone call from clinicians concerned about the “rising rates ” of trimethoprim resistance to E. coli…

Not being entirely convinced, I did a (20 year) search for E. coli resistance to trimethoprim at my lab, analysing over 2 million isolates, and came up with the following graph.

capture

 

I couldn’t work out how to insert a trendline into the graph (I am so useless…), but I think you will agree that it is going to be fairly flat.

The antibiotic apocalypse is not arriving in New Zealand anytime soon. In fact the whole concept of “antibiotic resistance” as perceived by the public is horribly generic and oversimplified…

This example above of course is just one microbe/antimicrobial combination out of many hundreds that could have been analysed, but the observation did highlight a couple of things to me:

  • If antibiotic usage is relatively constant in a population over a prolonged period of time, then antimicrobial resistance does not necessarily rise inexorably. (q.e.d.)
  • Always back your claims up with objective data wherever possible. It is the trends which are critical in the surveillance of antibiotic resistance. We are lucky that at my lab we can now search back through 20 years of electronic data. Before 1996 the data was paper based (and likely lost in a basement or incinerated by now!)

If you did a similar exercise for all the possible microbe/anti-microbial combinations (I just might if the Christmas holidays are quiet!), you will find some trends that are upwards, some that are static, and some where the resistance rate is trending downwards.

A bit like Twitter really….

So when someone says to you. “Antibiotic resistance is increasing all the time. In 10 years time, all infections will essentially be untreatable” (I really detest this type of generic, off the cuff, unsubstantiated statement…)

…you should respond with something along the lines of “Exactly which microbe and antimicrobial combination are you talking about?” and “Show me your data…”.

Some infections will be, and already are, untreatable (mostly due to extreme and focused selection pressure), but the chances of a whole bacterial species becoming pan-resistant are remote. There are two main reasons for this. i) Bacteria survive in open systems, and ii) Bacteria need to expend energy to become resistant.

But these are other stories altogether…

Michael