Chapter 9: Antimicrobial Drugs, Resistance & Chemotherapy

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Welcome to the Deep Dive, where we try to make complex topics clear and, well, fast.

Today we're tackling a really big one, antimicrobial chemotherapy.

It's something that truly changed the world, and central to this whole field is the term antibiotic.

Now, we use it all the time, but technically.

An antibiotic is usually a microbial product, something made by one microbe that can kill or stop the growth of other microbes.

Our plan today is to quickly walk you through the history.

Some of it's pretty wild.

Then the key ideas behind how these drugs work, how we test them, and finally tackle the huge challenge of resistance.

Think of it as a rapid rundown for anyone needing the core concepts fast.

Okay, let's dive in and maybe start somewhere unexpected.

The history of treating syphilis, this is kind of a bizarre story, this disease, treponema pallidum, it was awful, started with a painless sore, a chancre, but could lead to terrible neurological damage later on, or a syphilis, they called it general paresis of the insane.

Yeah, and for ages, the cures were just brutal.

There was this guaiac potion that basically made people vomit and sweat profusely, or mercury ointments.

Think about that, rubbing mercury onto people.

Mercury?

Seriously?

Oh yeah, it was incredibly toxic, sometimes killed the patient faster than the syphilis.

But horrifyingly, it was considered the best shot they had for a long time, real desperation.

Wow.

So the first glimmer of something more targeted came from Paul Ehrlich.

He got the Nobel Prize in 1908.

That's right.

He developed salversin, which was based on arsenic.

Now arsenic is still poison, obviously, but it was a specific chemical designed to target the spirachae.

It was a huge conceptual leap, even if the drug itself was rough.

A step up from just smearing mercury around.

But even arsenic wasn't the end of the weird treatments, was it?

Especially for that late -stage neurocyphilis.

No, not at all.

If salversin didn't work, some doctors turned to something called pyrotherapy.

The idea, believe it or not, was to induce a really high fever to, well, cook the syphilis bacteria to death inside the patient.

Cook them to death?

How?

Get this, they infected the patient with malaria.

You're kidding.

An Austrian doctor, Julius Wagner -Zharag, noticed malaria gives you these predictable high fevers.

And crucially, malaria could be treated afterwards with quinine.

So he'd intentionally give syphilis patients malaria, let the fever rage, and then cure the malaria.

And this worked.

Often enough that he won the Nobel Prize for it in 1927.

It just shows how desperate the situation was before we had truly effective selective drugs.

Giving someone malaria seemed like a reasonable option.

Okay, so moving away from intentionally giving people malaria,

what was the breakthrough principle that finally led to modern drug development?

It really comes back to Erlich's core idea, selective toxicity.

It's the absolute foundation.

The goal is the magic bullet.

A drug that kills the pathogen, the microbe causing the disease, but leaves our own human cells unharmed.

That makes sense, like a sniper rifle instead of a bomb.

Exactly.

And we measure how well a drug achieves this with a therapeutic index.

It's pretty simple, actually.

It's the ratio of the toxic dose, the amount that starts harming the patient to the therapeutic dose, the amount needed to actually treat the infection.

So a bigger number is better.

Absolutely.

A large therapeutic index means there's a wide gap between the dose that works and the dose that hurts.

That means fewer side effects, a safer drug overall.

Got it.

Now, when we talk about these drugs, there are different ways to classify them, right?

Like where they come from.

Yeah, you can look at the source.

Is it natural, like penicillin from a mold?

Is it purely synthetic, made in the lab like sulfa drugs?

Or is it semi -synthetic, where scientists take a natural antibiotic and tweak it chemically, like making ampicillin from penicillin?

And then there's the spectrum, what kinds of microbes they hit.

Narrow -spectrum drugs only work against a few types of bacteria.

Maybe just gram -positives, for example.

Broad -spectrum drugs hit a much wider range, gram -positives and gram -negatives.

And maybe the most important classification in practice is the effect, right?

Whether it kills or just stops growth.

Definitely.

We distinguish between cytol agents, those that actively kill the pathogen, and static agents, which just inhibit growth reversibly.

They basically pause the bacteria.

And why is that distinction so critical?

Well, static agents rely heavily on the patient's own immune system to come in and clear out those paused bacteria.

If someone is immunocompromised, their immune system might not be up to the task.

And a static drug might not be enough.

You might need something cytol to actually eliminate the infection.

So how do labs figure out if a drug is cytol or static, and how much is needed?

They measure two key things.

First,

the minimal inhibitory concentration, or MIC.

That's the lowest concentration of the drug that stops the bacteria from visibly growing in the lab.

You can't see any cloudiness in the test tube.

Okay, stops growth.

But that doesn't mean they're dead.

Exactly.

That's where the minimal lethal concentration, or MLC, comes in.

This is the lowest concentration that actually kills 99 .9 % of the bacteria.

Ah, so if the MLC is really close to the MIC.

Then you've likely got a cytol drug.

If the MLC is way higher than the MIC, or maybe you can't even reach it without toxic levels, then it's acting as a static agent.

And getting these MIC and MLC values right is obviously super important for treatment.

I assume there are standard ways labs do this.

Oh, absolutely.

Standardization is key for reliable results.

They use specific growth media, usually Mueller -Hinton -Broth or Agar, so results are comparable between labs.

So what does the test actually look like?

Well, the classic way to find the precise MIC and MLC is the dilution susceptibility test.

Picture a row of test tubes.

Each tube has the growth broth, but with successively lower concentrations of the antibiotic, like a dilution series.

Then you add the same amount of bacteria to each tube and incubate them.

Afterwards, you just look for the last tube in the series that has no visible plaudiness, no growth.

The concentration in that tube is your MIC.

Simple enough.

And the MLC.

To find the MLC, you take a little sample from the clear tubes, the ones at or above the MIC, and you spread it onto a fresh Agar plate that has no antibiotic.

You incubate that plate.

If nothing grows, it means the bacteria in the original tube were actually killed by that concentration.

That's your MLC.

Right.

But in a busy hospital lab, they often use something faster, visually, right?

The disc thing.

Yeah.

The disc diffusion method, also called the Courtney Bauer test, it's very common.

You take an Agar plate and spread the patient's bacteria evenly across the surface, like seeding a lawn.

Then you place these little paper discs on the Agar.

Each disc is impregnated with a known amount of a specific antibiotic.

And the antibiotic spreads out.

Exactly.

It diffuses out from the disc into the Agar, creating a concentration gradient.

It's highest right next to the disc and gets lower the further away you go.

So if the bacteria are susceptible...

They won't be able to grow near the disc where the concentration is high enough.

This creates a clear circle around the disc where the bacteria haven't grown.

That's called the zone of inhibition.

And the edge of that zone tells you something.

It tells you the MIC.

The very edge of that clear zone represents the point where the antibiotic concentration has dropped just low enough to allow growth.

So essentially, it marks the MIC for that drug against that bacteria.

So bigger zone means more effective drug.

Not necessarily.

This is a key point.

You measure the diameter of the zone, and labs have standardized charts that correlate that diameter to whether the bacteria is susceptible, intermediate, or resistant to that drug.

But you can't just compare zone sizes between, say, penicillin and tetracycline and say the with the bigger zone is better.

Why not?

Because the size of the zone also depends on how well the drug diffuses through the agar.

Its solubility and molecular weight matter.

A drug might make a huge zone just because it sprints easily, not because it's inherently more potent at the MIC.

Ah, okay.

That makes sense.

Is there a way to get a direct MIC reading from a pleat method?

Yes, there is.

It's called the A -test.

It's quite clever.

It's a plastic strip, and on one side it has a predefined continuous gradient of antibiotic concentration printed along its length.

So high concentration at one end, low at the other.

Exactly.

You lay this strip on the inoculated agar plate.

As the antibiotic diffuses out, it forms an elliptical zone of inhibition.

Where the edge of that ellipse crosses the strip, you can read the MIC value directly off the printed scale on the strip.

It gives you a quantitative MIC, like the broth dilution, but on an agar plate.

Very neat.

Okay, now for the really cool part.

How do these drugs actually work?

How do they pull off that selective toxicity?

It all comes down to targeting things that bacteria have, but we don't.

Or things that are significantly different between bacteria and human cells.

Like the cell wall.

I remember that's unique to bacteria.

Precisely.

Target one.

Cell wall synthesis.

This is often the best target, because human cells don't have peptidoglycan walls.

This gives these drugs a very high therapeutic index.

They're generally very safe for us.

And penicillins are the classic example here.

Penicillins and cephalosporins, yes.

They both have a key structural feature called the bollactam ring.

What they do is inhibit the final step of building the peptidoglycan wall, called transpeptidation.

This is where the wall components get cross -linked together to make it strong.

How do they inhibit it?

They bind to the enzymes that do the cross -linking.

These enzymes are called penicillin binding proteins, or PBPs.

By binding to the PBPs, the bollactam drugs block their function.

The wall can't be built properly, it becomes weak, and the bacterium often just bursts due to osmotic pressure.

Okay, so they hit the enzyme.

What about vancomycin?

That's another cell wall drug, right, used for serious infections.

Right.

Vancomycin is a big molecule, a glycopeptide.

It also blocks transpeptidation, but in a totally different way.

Instead of binding the enzyme, vancomycin binds directly to the building block itself specifically, to the Diala -Diala sequence at the end of the peptide chain that's about to be cross -linked.

So it sort of covers up the spot where the enzyme needs to work?

Exactly.

It physically obstructs the enzyme by binding to its substrate.

Because it's so large, it mainly works against gram -positive bacteria, which have that thick exposed piptidoglycan layer.

It's often a drug of last resort for things like MRSA.

Okay, target one, cell wall.

What's next?

Target two, protein synthesis.

This relies on differences between bacterial and human ribosomes.

Bacteria have 70S ribosomes, while our cells have 80S ribosomes in the cytoplasm.

They're different enough to target.

But you said earlier our mitochondria have 70S ribosomes, too.

That's the catch.

They do.

So while there is selective toxicity, the therapeutic index for protein synthesis inhibitors is generally lower than for cell wall inhibitors.

There's more potential for side effects, because they can sometimes interfere with mitochondrial protein synthesis.

Makes sense.

So what are some examples here?

Well, there are the imidoglycosides, like streptomycin.

They bind to the 30S subunit of the bacterial ribosome and cause it to misread the mRNA code or just stop translation too early.

They're bactericidal, killing the bacteria, but they can have serious toxicity, hearing loss, kidney damage.

It's a significant risk.

Tetracyclines also target the 30S subunit, but they're generally bacteriostatic.

They stop growth.

They're broad spectrum.

Then you have macrolides, like erythromycin, which bind to the 50S subunit and block the ribosome from moving along the mRNA.

They're often used if someone has a penicillin allergy.

Are there newer ones?

Yes.

A really important newer class is the oxazolidinones, like linazolid.

These are synthetic and have a unique mechanism.

They bind to the 50S subunit, but prevent the whole 70S ribosome from even assembling correctly at the start of protein synthesis.

They're reserved for tough, resistant gram -positive infections like MRSA and VRE.

Okay, cell wall, protein synthesis.

What about hitting their metabolism?

Target three, metabolic pathways.

These are often called anti -metabolites.

The classic example is exploiting the folic acid pathway.

Because we get folate from our diet, but bacteria make their own.

Exactly.

That difference is key for selective toxicity.

The sulfonamides, or sulfa drugs,

were among the very first antibiotics discovered.

They look structurally very similar to a molecule called paeba, which is needed for the first enzyme in the bacterial folic acid synthesis pathway.

So they trick the enzyme.

Pretty much.

They act as competitive inhibitors.

They bind to the enzyme instead of paeba, blocking the pathway.

No folate synthesis, no growth.

And there's another drug that hits the same pathway.

Yes, trimethoprim.

It's also synthetic,

and it inhibits a different enzyme, one acting later in the same folic acid pathway.

And using them together is better.

Much better.

This is a great example of synergistic drug interaction.

Blocking the pathway at two different points with both a sulfa drug and trimethoprim is far more effective than using either drug alone.

That combination is used very commonly.

Clever.

Okay, one more main target group.

Target four, nucleic acid synthesis, disrupting DNA replication or RNA transcription.

How do they do that selectively?

Well, the fluoroquinolones, like ciprofloxacin, are synthetic drugs that inhibit bacterial enzymes called topoisomerases, specifically DNA gyrase in many bacteria.

These enzymes were essential for managing the supercoiling and uncoiling of DNA during replication and repair.

Bacterial topoisomerases are different enough from ours for selective targeting.

Are they safe?

They are bactericidal and broad spectrum, but there are growing safety concerns.

They carry warnings about potentially disabling side effects like tendon rupture or nerve damage so their use is becoming more restricted.

Okay.

What about hitting RNA?

That's where the rifamysins, like rifampin, come in.

They specifically bind to the beta subunit of the bacterial RNA polymerase, the enzyme that makes RNA from a DNA template.

This blocks transcription.

Rifampin is really important, especially in multi -drug treatments for things like tuberculosis.

Wow.

Okay, so we've covered bacteria pretty well, but what about treating things like fungi or protozoa?

You mentioned that's harder.

Yeah, it connects right back to that selective toxicity problem.

Fungi and protozoa are eukaryotes, just like us.

Our cells share many more similarities with them than with bacteria.

That means fewer unique targets, which translates to a lower therapeutic index and often more side effects with the drugs we do have.

So for fungi, what's the main target?

The best unique target we have in fungi is their cell membrane.

Fungal membranes contain a sterol called ergosterol, whereas our cell membranes primarily use cholesterol.

That difference is exploitable.

There are two main approaches.

The polyenies, like amphotericin B and nystatin, bind directly to the ergosterol in the fungal membrane.

This disrupts the membrane structure, makes it leaky, and kills the cell.

Amphotericin B, I've heard that one's rough.

Extremely.

It's often nicknamed amphoterable because of its significant toxicity, especially kidney damage.

But for severe, life -threatening systemic fungal infections, sometimes it's the only option despite the risks.

So it's a tough balancing act.

Definitely.

A somewhat safer approach involves the azoles, like gluconazole.

Instead of binding to existing ergosterol, they block the synthesis of ergosterol.

Less ergosterol means a faulty membrane.

They tend to be better tolerated than amphotericin Bs.

Any others for fungi?

There's also 5 -flucytosine.

It's interesting because it gets converted into an active toxic form by fungal enzymes, but not typically by human enzymes.

This active form messes with fungal RNA function.

Okay, moving to protozoa.

Malaria is a huge one, right?

Plasmodium.

Yes.

Traditionally, quinine and its derivatives, like chloroquine, were mainstays.

They work by interfering with the parasite's ability to detoxify heme, a breakdown product of the hemoglobin it consumes from red blood cells.

The parasite normally polymerizes toxic heme into nontoxic hemozoin, and chloroquine blocks this process.

But there's resistance now.

Widespread resistance to chloroquine, unfortunately.

So now, the standard is combination therapy, often including artemis and derivatives, which come from the sweet wormwood plant.

These combinations hit the parasite in multiple ways.

What about other protozoa, like amoebas?

For anaerobic protozoa, like entamoeba histolytica, which causes amoebic dysentery, or giardia, a drug called metronidazole is very effective.

It gets activated under anaerobic conditions into a reactive form that damages the parasite's DNA.

And you mentioned something about an apicoplast.

Right.

Some protozoa, including the malaria parasite, plasmodium, and toxoplasma, belong to a group called apicomplexans.

They have this unique organelle called an apicoplast, which seems to be derived from an ancient chloroplast.

Like in plants.

Sort of, yeah.

And importantly, this apicoplast has its own 70S ribosomes, just like bacteria.

So some antibiotics that target bacterial protein synthesis, like certain aminoglycosides or tetracyclines, can actually inhibit protein synthesis within the apicoplast, harming the parasite.

It's a fascinating example of finding a unique internal target.

Very cool.

Okay, last group.

Viruses.

You said they aren't technically antibiotics.

Correct.

We call them antivirals.

They don't target living cells in the same way.

Viruses hijack our own cell machinery, so antivirals have to target specific steps in the viral life cycle.

Generally, they limit how long you're sick or how severe it gets, but very few offer a complete cure.

Like Tamiflu for the flu.

Exactly.

Oseltamivir inhibits a viral enzyme called neuraminidase.

Influenza viruses need this enzyme to cut themselves free from the host cell surface after they replicate.

Tamiflu blocks that release, so new viruses can't spread as effectively.

What about viruses like herpes?

For DNA viruses like herpes simplex or varicella zoster, chicken pox shingles, drugs like a cyclovir are common.

They're nucleoside analogs.

They look like the building blocks of DNA.

Here's the clever part.

They need to be activated, phosphorylated to work.

And the viral enzyme, thymidine kinase, is much better at doing this first phosphorylation step than our own enzyme.

So the virus activates its own poison.

Essentially, yes.

Once activated and incorporated into the growing viral DNA chain, a cyclovir lacks the proper chemical group for the next nucleotide to attach.

This causes chain termination DNA synthesis stops.

And HIV, that treatment seems really complex.

It is.

HIV is a retrovirus, and it mutates incredibly rapidly, leading to resistance.

So effective treatment absolutely requires a cocktail of drugs, usually at least three different agents hitting different targets simultaneously.

There are several classes.

Reverse transcriptase inhibitors blocking RNA to DNA, protease inhibitors blocking viral protein processing,

integrase inhibitors blocking viral DNA insertion into host genome, and adhesion inhibitors blocking entry into the cell.

Wow.

Hitting it from all sides.

Any recent breakthroughs?

Huge ones for hepatitis C, HCV.

We now have direct acting antiviral agents, DAAs, like Sophosbuvir.

These drugs directly target essential HCV enzymes, like its RNA polymerase.

Combination DAA therapy can now cure most HCV infections, which is a massive leap forward from older treatments.

That's incredible progress.

But despite all these amazing drugs, we have to talk about the flip side.

The growing crisis of antimicrobial resistance.

Yeah, it's arguably the biggest public health threat we face globally.

We're seeing resistant strains like MRSA, methicillin -resistant Staph .areus, VRE, vancomycin -resistant enterococci, CRE, carbapenem -resistant enterobacteriaceae.

These are nightmare bacteria.

And the projections are scary.

Terrifying.

Some reports predict that by 2050, drug -resistant infections could kill more people annually, like 10 million, than cancer does today.

It threatens to undo decades of medical progress.

So how does resistance even happen?

Is it just bad luck?

Well, microbial evolution is incredibly fast.

We first distinguish between intrinsic resistance, where a bacterium is just naturally not affected by a drug.

For example, mycoclasma species lack a cell wall entirely, so penicillin and other cell wall inhibitors just have no target.

They're intrinsically resistant.

OK, that's built in.

What's the dangerous kind?

Acquired resistance.

This is where a previously susceptible bacterium becomes resistant through a genetic change.

This can happen via a random mutation, or more commonly and rapidly, through horizontal gene transfer, getting resistance genes from other bacteria, often on plasmids or transposons.

And what do those genes actually do?

How do they make the bacteria resistant?

There are several main mechanisms.

Let's stick through them.

First, they can modify the target of the drug.

Like change the locks so the key doesn't fit.

Perfect analogy.

MRSA does this.

It acquires a gene, MEKA, that encodes a different penicillin -binding protein, PBP2A, that molactam antibiotics just don't bind too well.

The drug is there, but it can't hit its target effectively.

Similarly, VRE changes the vancomycin binding site on the cell wall precursor from DLADL to DLADLactate.

Vancomycin can't bind tightly anymore.

OK, change the target.

What else?

Second, they can inactivate or degrade the drug.

The absolute classic example here is volactamase enzymes,

including penicillinase.

Bacteria with genes for these enzymes produce proteins that literally chop open the volactam ring of penicillins and cephalosporins, destroying the antibiotic before it can even reach the PBP target.

So they have an antidote, basically.

In a way, yes.

And there are enzymes that modify or degrade other classes of antibiotics, too.

Third mechanism,

minimize the drug concentration inside the bacterial cell.

How do they do that?

Keep it up.

Sometimes by altering their outer membrane porins to reduce uptake, especially in gram negatives.

But a really major mechanism is using efflux pumps.

These are transport proteins embedded in the bacterial membrane that actively pump the antibiotic right back out of the cell, sometimes as fast as it gets in.

Like a bilge pump on a boat.

Exactly.

And what's particularly worrying is that many of these efflux pumps are quite nonspecific.

A single type of pump might be able to recognize and expel multiple different classes of antibiotics.

This is a major route to multidrug resistance, MDR.

That sounds really bad.

One pump defeats multiple drugs.

It is.

And the fourth main way is to develop an alternate pathway, or increase production of the target.

For example, some bacteria become resistant to sulfa drugs, not by changing the enzyme, but by figuring out how to scavenge preformed folic acid from their environment,

completely bypassing the need for the inhibited synthesis pathway.

So they just find a detour around the roadblock.

Precisely.

These mechanisms, often spread rapidly on mobile genetic elements, are why resistance is such a formidable challenge.

So what can we actually do about it?

It sounds kind of hopeless.

It's not hopeless, but it requires a massive coordinated effort.

Key strategies include much tighter control and stewardship of existing antibiotics, using them only when necessary.

Using the right drug at the right dose for the right duration.

We need to prevent overuse and misuse in both medicine and agriculture.

Finishing the full prescription is part of that, right?

Absolutely critical.

Stopping treatment early allows the more resistant survivors to multiply.

We also need to use drugs in sufficiently high concentrations when appropriate and rely more on drug combinations or cocktails, especially for infections known to develop resistance easily, like TB, HIV,

and sometimes malaria.

Hitting the bug with multiple mechanisms at once makes it much harder for resistance to emerge.

Are there new things on the horizon?

New types of drugs?

There's ongoing research, definitely.

Things like structure -based rational drug design aim to create drugs that are less susceptible to existing resistance mechanisms.

Scientists are looking for entirely new targets, like FTSC, a protein involved in bacterial cell division.

What about completely different approaches?

There's a renewed and growing interest in bacteriophage therapy.

Phages are viruses that specifically infect and kill bacteria.

The idea is to use phages that target specific drug -resistant strains.

It's an old idea, actually, that's getting a fresh look because of the resistance crisis.

Fascinating.

Okay, let's try to wrap this up.

Key takeaways from our deep dive today.

I'd say number one is the concept of selective toxicity,

Ehrlich's magic bullet that underpins the entire field.

Number two is the importance of standardized testing, like Kirby Bauer and MIC determination, for guiding treatment effectively.

And number three has to be the mechanisms understanding how drugs like penicillins target the cell wall or amnoglycosides hit ribosomes is key, but also understanding how bacteria fight back through target modification, drug inactivation, or efflux pumps that's crucial for appreciating the resistance challenge.

Right, and maybe that leads to a final thought to leave you with.

Think about the difference in safety between, say, penicillin, which hits the bacterial cell wall and has a huge therapeutic index, versus something like afotericin B used for fungal infections, which hits ergosterol but is notoriously toxic with a low therapeutic index.

Because fungi are eukaryotes, closer to us.

Exactly.

What does that huge difference in safety profiles tell you about the fundamental difficulty, the inherent challenge, of developing drugs against organisms that share our basic cellular structure?

And how should medicine constantly weigh the potential benefit of a cure against the very real risk of harm to the patient, especially when dealing with these more difficult to treat eukaryotic pathogens?

That is a heavy question, balancing the magic bullet against potential collateral damage.

Definitely something to think about.

Thank you so much for walking us through all of this.

My pleasure.

It's a critical topic.

And thank you for joining us on the Deep Dive for this look at antimicrobial chemotherapy.

We hope it was clear and helpful.