When Water Runs Short: A 5-Point Decision Guide for Decontamination Protocols

You're in a field clinic. Ten patients, maybe more. Water runs from a tank that's half empty and won't be refilled for two days. Standards say you should wash surfaces with soap and water before disinfecting—but standards weren't written for this. So what do you do?

Across outbreak responses—cholera, Ebola, COVID-19—water scarcity twists every decision. Protocols that work in a hospital sink fail when you're hauling buckets. This isn't a minor inconvenience; it's the difference between breaking a transmission chain and watching it persist. Here's a 5-point guide built for the real constraints of field operations.

Why Water Scarcity Is the Default in Outbreak Response

When the Tap Runs Dry: The Real Starting Point

Most decontamination protocols you find in textbooks were written for places with reliable water supplies—hospital basements with endless tap pressure, treatment plants with tanker trucks on standby. That sounds fine until you're standing in a cholera treatment center in the back of a pickup, watching your last 50 liters of clean water get shared between staff handwashing and patient rehydration. I have seen that exact math fail. The hard truth is that water scarcity is not an exception in outbreak response—it's the rule. Logistics breakdowns, damaged infrastructure, or simply the sheer volume of disinfectant mixing required means you often have a fraction of the water the manual demands.

What usually breaks first is the assumption that you can rinse. Standard protocols lean heavily on copious water for pre-cleaning, dilution, and final rinsing. Without that water, a 0.5% chlorine solution leaves residue that corrodes surfaces, or worse, it fails to contact the pathogen because organic matter was never flushed away. The catch is that reducing water use doesn't mean reducing safety—it means matching the method to what you actually have on hand. Otherwise, you create a false sense of security: surfaces look clean, but the viral load remains.

How Water Shortages Amplify Infection Risks—Silently

Here is the pattern I have watched repeat itself in field settings. When water gets tight, teams start cutting corners. They reduce contact times because they're rationing the mix. They skip the second wipe-down because they have no rinse water. They reuse cloths. The odd part is—these are experienced responders, not novices. The shortage forces them into trade-offs they never planned for. And the pathogen doesn't care about their good intentions. A single skipped rinse on a high-touch surface in a treatment unit can seed a cross-contamination chain that shows up in case numbers three days later.

‘We ran out of water for the third rinse. We told ourselves it was fine. It was not fine. Six staff went down the next shift.’

— anonymous field report, MSF cholera response, 2019

The silence around this problem is dangerous. Most outbreak after-action reports focus on drug stockouts or laboratory delays, not the mundane fact that the water bladder had a pinhole leak at 2 a.m. But in practice, that leak can unravel the entire decontamination plan. Standard protocols assume abundant water. That assumption is a liability, not a luxury, in the places where outbreaks actually burn hottest.

Wrong starting point, every time.

The Core Idea: Match the Method to the Pathogen and Surface

Enveloped vs. Non-Enveloped Virus Susceptibility

The first cut in any decontamination decision is simple: is the virus wrapped in a lipid coat or not? Enveloped viruses—SARS-CoV-2, Ebola, influenza—are soft targets. A standard quat wipe or 0.05% bleach solution disrupts that fatty envelope in under sixty seconds. We fixed this by treating them all with the same low-concentration protocol during a Marburg scare in 2022; it worked. Non-enveloped viruses—norovirus, adenovirus, hepatitis A—are stripped down, tough. They laugh at quats. You need higher chlorine doses (≥2,000 ppm) or accelerated hydrogen peroxide to crack their protein shell. That difference alone determines whether your team uses a spray bottle or a full bucket soak. The catch: most outbreak field kits stock only one broad-spectrum disinfectant. If you guess wrong on the envelope, you waste time and water.

Wrong order. Check the lab report before you mix.

Spore-Formers: The Toughest Challenge

Clostridium difficile. Bacillus anthracis. Spore-formers store their genetic material in a crystal-like coat that survives boiling, drying, and most disinfectants. I have seen field teams hit anthrax spores with 10% bleach for ten minutes—and still get regrowth on contact plates. That hurts. For spore-formers, you need a sporicide: peracetic acid (≥0.2%) or chlorine dioxide gas. Neither is common in a standard cholera kit. The trade-off is brutal—sporicides corrode metal fittings, burn skin on contact, and require precise ventilation. Most guidelines skip this edge case entirely. The odd part is: if you're treating a cholera bed mat, you never face spores. But open a suspicious powder on that same cot, and the protocol flips upside down. Surface matters, but the pathogen’s resistance tier matters more.

Reality check: name the epidemiology owner or stop.

Reality check: name the epidemiology owner or stop.

‘A spore is not a germ. It's a survival capsule. Treat it like a bomb, not a spill.’

— field trainer, Médecins Sans Frontières, 2019

Surface Material Effects on Disinfectant Performance

Match the method to the material, or the method fails. Porous surfaces—wood, unfinished concrete, fabric—absorb liquid. A 500-ppm chlorine spray on a wooden bench delivers maybe 100 ppm at the surface; the rest wicks into the grain. Most teams skip this: they spray, watch it pool, assume coverage, and leave. The seam blows out when a child touches that bench an hour later. Non-porous surfaces—stainless steel, glass, smooth plastic—hold the disinfectant film, but they also let it evaporate faster if the ambient air is dry. That's a water-scarcity paradox: in a dry outbreak zone, your chemical sits on steel for ninety seconds and leaves a crystal residue instead of a wet kill time. What usually breaks first is the contact time, not the chemical strength. We fixed this by switching to wipes pre-wetted with a humectant (propylene glycol) that slows evaporation by 30%. Not everyone can source that. The call is: for porous surfaces, use a longer soak and double the volume. For non-porous, cut the volume but guarantee the clock. One rhetorical question worth asking: how many outbreaks have been prolonged because a team disinfected the wrong surface type with the right chemical?

You lose a day. Returns spike.

How the Key Disinfectants Work Without Much Water

Alcohol-Based Hand Rubs and Surface Wipes: Speed vs. Soil

Ethanol at 70–80% concentration kills most enveloped viruses and vegetative bacteria in under thirty seconds. That speed is its superpower when water is measured by the liter, not the hour. You spray, you wipe, you move on. But the catch—and it’s a hard one—is organic matter. Blood, vomit, or even a thin layer of dried mud on a surface will shield pathogens from the alcohol. I have watched teams waste minutes scrubbing a visibly dirty tabletop with an ethanol wipe, only to see their rapid test still light up positive. The alcohol denatures proteins, yes, but it can't penetrate a biofilm or a crust of field soil. So the rule is simple: alcohol works brilliantly only on pre-cleaned surfaces. That means you still need water for the first pass—or you need to accept that alcohol wipes are a finishing step, not a primary decon tool for gross contamination. Also, the stuff evaporates fast, especially under a tropical sun; contact time can drop below the required threshold if you wipe and walk away. Wring out your wipe slowly, keep the surface visibly wet for the full dwell time—that hurts compliance but it works.

Chlorine-Based Solutions: The Workhorse That Thirsts

Chlorine compounds remain the backbone of field disinfection because they're cheap, broad-spectrum, and relatively forgiving of operator error. Sodium hypochlorite (household bleach) at 0.5% solution kills bacteria, viruses, and most fungi within ten minutes—provided the solution is fresh. That last part is the pitfall. Mixed chlorine degrades in direct sunlight and in warm containers; after six hours in a jerry can left on the ground, the effective concentration can drop below 200 ppm. The field fix is to prepare small batches, use opaque carboys, and test with a chlorine meter every shift. But here is the real water problem: a standard chlorine dip or spray uses roughly 1–2 liters per square meter of surface. In a cholera treatment center, that adds up to hundreds of liters a day. You can stretch it by using a trigger-sprayer instead of a bucket-and-rag—that cuts water use by about 60%—but you lose mechanical scrubbing action. The emulsion between chlorine and organic material also consumes free chlorine fast; for blood spills you need 5,000 ppm, which means even more concentrated stock and even less water efficiency. So the trade-off is volume versus safety.

“Chlorine works miracles—until the bucket runs dry at noon and the next resupply is three hours out.”

— Field note from a MSF logistics officer, 2022

Quaternary Ammonium Compounds: Low-Water, High-Caution

Quats, as they're called, are surface-active disinfectants that cling to pathogens and disrupt their membranes. The biggest draw for water-scarce settings: many quat formulations require no rinse. You spray, let it sit for the labeled contact time (usually five to ten minutes), and walk away. No bucket, no wiping dry, no residual water on the floor. That saves liters per shift. But the limitations stack fast. Quats have poor activity against non-enveloped viruses—norovirus, polio, rotavirus—which means they're useless for most outbreak contexts unless you verify the pathogen first. They also get inactivated by cotton fibers, so using a quat wipe on a cotton bandage? Wrong order. And hard water with high calcium or magnesium can precipitate the quat molecules, dropping efficacy to zero. I have seen teams deploy quat sprays in a field clinic only to realize later that the local well water, untreated, rendered every spray bottle effectively inert. You need buffered, low-hardness water for the mix—which adds a pre-treatment step that many emergency protocols skip. So quats are a good option for low-risk surfaces in an aid station, but not for a confirmed outbreak ward.

Hydrogen Peroxide Vapor and UV-C: Advanced But Fragile

These are the heavy hitters when water is truly gone—zero liters for liquid decontamination. Hydrogen peroxide vapor (HPV) uses a dry fog or vaporized 35% solution that condenses on surfaces and releases reactive oxygen species. No rinse, no wipe, no water. Contact time is short—about fifteen minutes to kill anthrax spores on a clean surface. But the machine is expensive, requires power, and the vapor is toxic during the cycle: you seal the room, evacuate people, run the cycle, then ventilate for another thirty minutes. Not a tool for a triage tent with patients streaming in. UV-C light works similarly—physically disrupts DNA—but it casts shadows, so vertical surfaces, undersides of tables, and corners remain contaminated unless you use multiple fixtures or a robot. The odd part is that UV-C lamps lose output over time; a fixture that ran 8,000 hours may be emitting only half its labeled dose. No one in the field carries a radiometer. So these options are for terminal disinfection of a closed room or an ambulance after patient transfer. Not your daily surface protocol. Not your outbreak quick fix.

A Walkthrough: Choosing a Protocol for a Cholera Treatment Center

‘We burned through our 500-liter reserve in three hours. After that, every drop had to count.’

— field log, MSF cholera response, 2022

That blunt reality sets the stage for this walkthrough. A cholera treatment center (CTC) in a water-scarce zone. The pathogen: Vibrio cholerae, gram-negative, fragile outside the host—enveloped, yes, but shed in massive volumes. Diarrheal fluid, soiled bedding, latrine floors, patient hands, reusable rubber boots. Each surface demands a different contact time. Each requires a different water volume. The decision tree starts with a single question: can you spare even one liter per patient bed, per shift? Most teams answer no. So you default to a 0.05% chlorine solution—sodium hypochlorite, 500 ppm free residual chlorine, prepared fresh every four hours. That's your workhorse. Why? Because chlorine degrades fast in tropical heat, but when water is tight, you need a disinfectant that works within a five-minute wet time on hard, non-porous surfaces. Alcohol-based disinfectants (typically 70% ethanol or isopropanol) demand 30–60 seconds of contact, but they evaporate completely—zero water footprint. The trade-off is brutal: alcohol is wasted on porous surfaces like untreated wood or cracked concrete. The seam blows out. You lose a day.

The tricky bit is matching the method to the moment. Inside a CTC, you have three surface categories: patient-care surfaces (plastic mattresses, IV poles, examination tables), high-touch non-critical items (door handles, tap handles, shared stethoscopes), and soiled zones (latrine seats, vomit buckets, floor areas within one meter of a bed). For the soiled zones, a 0.05% chlorine soak—five minutes wet, then rinse with the minimal water you have. That hurts. I have seen teams skip the rinse step to save water. Don’t. Residual chlorine corrodes metal and irritates skin; in a cholera ward, irritated skin on caregivers’ hands leads to glove breaches, which leads to staff infection. Most teams skip this: the rinse water can be captured in a drum and used for the next chlorine dilution. Closed-loop thinking. Not elegant, but field-hardy.

Flag this for epidemiology: shortcuts cost a day.

Flag this for epidemiology: shortcuts cost a day.

Comparing 0.05% chlorine solution vs. alcohol-based disinfectants

Here is where the decision splits. On clean, smooth surfaces—stainless steel, glazed tile, laminated countertops—alcohol wins on speed and zero water demand. But alcohol is flammable. You can't store forty liters of 70% ethanol in a tent at 40°C. I watched a colleague try. The bottle caps swelled, the seals popped, and we lost half the stock to evaporation. Chlorine, mixed as a 0.05% solution from powder or tablets, is safer in bulk but requires fresh water for dilution and rinse. The calculus: one liter of alcohol wipes roughly 20 square meters of surface. One liter of chlorine solution wipes the same area but requires another 0.3 liters of rinse water. Over 200 patient beds per day, that difference compounds fast. When water trucks arrive only every 48 hours, alcohol is the friend of the night shift—used sparingly on stethoscopes, phone screens, and laryngoscope handles. Chlorine is for the gross mess. Wrong order kills people: if you deploy alcohol on a stool-smeared floor, the organic load neutralizes it instantly. Clean first, then disinfect. That sequence matters more than which chemical you pick.

Step-by-step decision with water constraints, contact time, and safety

Let me walk you through a real shift. 0600 hours: water tank reads 300 liters, expected resupply at 1800. You have twelve hours to run the CTC. First, allocate 200 liters for handwashing stations—that's non-negotiable, cholera is a fecal-oral disease, staff hands are the transmission highway. You're left with 100 liters for all disinfection duties. That's thin. A single chlorine mop bucket holds 10 liters; at 0.05% concentration, you need to change the solution every two hours or when visibly soiled—whichever comes first. So you have ten bucket-refills for the entire day. Divide by three shifts: three buckets per shift, plus one emergency bucket. The math forces a decision: reduce surface area or switch to alcohol for non-soiled zones. We fixed this by designating a ‘low-touch zone’ map taped to the supply tent—door handles, light switches, and chair backs get alcohol wipes. Everything within splash range gets chlorine. The contact time for chlorine on smooth, pre-cleaned surfaces is five minutes. On porous surfaces like woven plastic tarps, ten minutes. That extra five minutes per patient bay adds up. One trick: start the clock when you begin wiping, not when you finish. Most teams wait until the whole bay is wet. That mistake cuts effective contact time in half. Set a timer. No exceptions.

The catch is safety. Chlorine fumes at 0.05% are tolerable for ten minutes, but by the third bucket refill in a closed tent, respiratory irritation kicks in. Ventilation matters. If the CTC is a military tent with no cross-breeze, you rotate staff every twenty minutes. Alcohol fumes are less irritating but flammable—no open flames, no smoking, no battery-charging stations near the disinfection area. I have seen a phone charger spark ignite alcohol vapor on a folding table. The fire was out in four seconds, but the panic cost us forty minutes of treatment time. The final decision point is the easiest: if the patient-to-staff ratio exceeds 10:1, forget any protocol that requires individual wipe-downs. Go to spray-and-wipe with chlorine, accept the higher water use, and train one designated disinfection staff member per shift. That person owns the bucket, the timer, and the log sheet. No delegation. No shortcuts. When water runs short, the protocol that survives is the one with a single, tired person executing it in the dark at 2 AM. Build for that person.

Edge Cases: Blood Spills, Food Surfaces, and Freezing Temps

Blood Spills: When Standard Dilution Kills Too Slowly

The protocol sheets say 'follow manufacturer instructions.' In a field tent, that page is gone—and the spill is spreading. Bloodborne pathogens change the math entirely. Hepatitis B and C, HIV—these are not your average vegetative bacteria. They demand a higher free-chlorine concentration, often 5000 ppm instead of the usual 1000–2000. And contact time? Thirty minutes minimum, not the five you might use for a dirty floor. I have seen teams grab the same spray bottle they use for general disinfection and hose down a bloody mattress. That's a protocol failure dressed up as speed. The catch is that a strong chlorine solution at that level corrodes metal in hours and bleaches fabric permanently—ruin a stretcher, and you're down a stretcher. For porous surfaces like unpainted wood or cracked plastic, no low-water method reliably penetrates the crevices. The honest fix: remove the contaminated material entirely if you can. If you can't, triple the volume and double the scrub time. That hurts resupply, but it beats a secondary outbreak.

Food Surfaces: Toxicity Isn't a Suggestion

You don't spray bleach on a plate and hand it back to a patient. Obvious, right? Yet I have watched exhausted staff mist chlorine onto cutting boards used for therapeutic feeding. The residue stays. Chlorine compounds, especially at field-strength concentrations, leave chemical byproducts that can cause nausea or worse in malnourished individuals. For food-contact surfaces in a water-short setting, thermal treatment is your only reliable friend—boiling water, steam (if you have a pressure cooker and fuel), or dry heat for metal tools. Alcohol-based wipes work on stainless steel utensils if the contact time is respected—thirty seconds wet, left to air dry—but alcohol evaporates fast in wind and is a fire risk near cookstoves. The trade-off is simple: you trade water for fuel. That means more smoke, more cooking time, more logistics. Most teams skip this: they rinse with chlorinated water and call it clean. Residue lingers. Test it with a simple pH strip—if the surface reads above 8 after drying, it's not safe for direct food contact. Wash again with boiled water, even if it's only a cup.

Freezing Temps: Where Alcohol Freezes and Chlorine Fails

Cold is the silent protocol-killer. Standard ethanol-based hand rubs and surface wipes freeze at around -10°C (14°F). You keep a bottle in your jacket pocket—it stays liquid. The second you put it on a cold table, it slushes. In a subzero cholera camp, the alcohol gels separate into icy sludge and useless water. That leaves chlorine. But chlorine's disinfection power drops sharply below 10°C. The reaction slows. Contact times quoted for 20°C may need to be tripled at 0°C—if the solution is still liquid. The odd part is—frozen surfaces look clean. Ice can trap pathogens in a suspended state. Once thawed, the bacteria resume activity. The fix is not elegant: pre-warm the disinfectant solution to 30–35°C before application. That requires a heat source and a thermos. Or move to a quaternary ammonium compound, which stays active at lower temperatures—but it's slower and has its own residue issues. Nothing works perfectly in frozen conditions. We fixed this once by rotating two spray bottles: one in use, one under a coat against a warm torso. Crude. But the outbreak didn't pause for the thermometer.

‘The worst mistake is assuming any single method survives the field intact. Cold, blood, and food each break the rulebook differently.’

— field coordinator, after losing a day to refrozen alcohol stocks

That said, you can test your protocols before deploying. Take a thermometer and a stopwatch into a cold room. Spray your chosen disinfectant at the target temperature. Measure the wet contact time. If it dries in under two minutes, or if the liquid is slush, the protocol needs redesign. Don't wait for a spill to discover the flaw.

Where This Approach Hits Its Limits

Inability to remove organic matter without water

Here is the brutal truth about low-water disinfection: it can't clean. You can spray, fog, or wipe with high-concentration chemistry until your gloves rot, but if that surface is caked with vomit, blood, or soil, the disinfectant will hit a protein wall and stop. Pathogens hide inside that organic load. I have seen teams in a Cholera Treatment Center spray a chlorine solution over a visibly stained mattress — and call it 'done.' The next patient got infected. Without water to flush, scrub, and lift debris, you're just wetting the dirt. The catch is that soap-and-water scrubbing uses ten to twenty times the volume of water that a typical low-water protocol allows. You trade microbial safety for logistical survival. That trade holds until someone dies because of it.

Wrong order. Some teams reverse the sequence — disinfect first, then try to clean. That fails too. Organic matter neutralizes chlorine, quats, and peroxides within seconds. You lose half your active chemical before it touches a single virus particle. Most field manuals skip this because the math is ugly: no water, no cleaning. No cleaning, no reliable kill.

Corrosion and damage to equipment

What usually breaks first is not the protocol — it’s the gear. Chlorine dioxide and sodium hypochlorite at the concentrations needed for low-water work (often 2000–5000 ppm) eat through metals like acid through paper. I have watched a brand-new sprayer’s brass nozzle pit and seize in three days. Collapsible water bladders develop pinhole leaks where concentrate pooled. Even stainless steel tables in a field hospital begin to bloom rust after two weeks of daily fogging with peracetic acid. The odd part is — manufacturers rarely mention corrosion in their glossy 'low-water solution' brochures. They test for thirty minutes in a lab, not thirty days in a monsoon. We fixed this by switching to PVC and HDPE fittings, but that meant carrying three different types of nozzles and knowing exactly which chemical was in which tank. One mistake, and the entire spray system seizes up mid-outbreak.

Not yet. Many groups try to dilute the chemical to protect equipment. That undercuts the contact time. You preserve the gear but lose the kill. The real constraint is material compatibility tables — those are rarely printed on waterproof paper. They disintegrate in the first rain.

Odd bit about epidemiology: the dull step fails first.

Odd bit about epidemiology: the dull step fails first.

Training and compliance challenges

You can design the perfect low-water protocol on paper. In practice, it requires every single staff member — nurses, cleaners, drivers — to follow a specific sequence under stress. Miss one step: don the respirator before mixing. Skip the pre-wet dwell time. Fail to rotate spray patterns. That protocol collapses. I have seen trained clinicians pour a pre-mixed disinfectant into an empty bottle for 'convenience,' then forget which bottle held which concentration. The result: a surface doused with plain water, not disinfectant, while the team believed they had completed a full decontamination cycle. The human factor is not a footnote. It's the limiting variable that turns a perfect algorithm into a perfect hazard.

‘The protocol works until the person holding the sprayer is exhausted, hungry, and scared.’

— field logistics coordinator, MSF cholera response, 2023

Training takes time. Outbreaks don't wait. We tried checklists laminated to sprayer handles — they got lost or splattered. We tried color-coded caps for concentrates — staff swapped them between shifts. What finally worked was a single-page visual guide (only pictures, no text) plus one supervisor whose only job was to walk the line and correct sequence errors. Even then, compliance hovered around seventy percent. The missing thirty percent is where outbreaks get second winds.

Reader FAQ: Your Most Pressing Field Questions

Can I reuse disinfectant wipes?

Short answer: no. The longer, field-hardened answer: it depends on the wipe and the soil load, but you should treat every used wipe as spent. Once a wipe touches a surface, it picks up organic matter—blood, vomit, dust—that neutralizes the active chemical. That same wipe, dipped back into the bucket, contaminates the whole solution. I have seen teams try to stretch a box of wipes across a full shift. The result? A pinkish film that smells clean but fails the swab test every time. The catch is cost per unit—good wipes are expensive—but a reused wipe is a false economy. If you must ration, cut wipes in half before use, never after.

Wrong move: dunking a used wipe into a fresh disinfectant bottle. You just seeded the bottle.

What if I only have clear liquor?

That bottle of high-proof spirits in your pack—vodka, moonshine, medicinal alcohol—feels like a solution. It isn't. Most clear liquors sit at 40% ethanol (80 proof). Effective disinfection requires 60–80% ethanol. Below that threshold, the alcohol can't denature proteins fast enough. The odd part is—higher is not better either. Pure ethanol (95%) evaporates too quickly, leaving no contact time. So that 120-proof rum? Useless against norovirus, poor against bacteria. What it can do: flush a small wound if nothing else exists. But for surface decontamination in an outbreak zone? Leave it for morale, not protocols.

One exception I have seen accepted in the field: 70% isopropyl alcohol poured over a clean rag, used once, then discarded. That works. Your uncle's homemade whiskey doesn't.

How long does chlorine stay active in sunlight?

Not long. Minutes. Direct UV radiation breaks down free chlorine fast—half-life can drop below 30 minutes in tropical midday sun. That means your bucket of 0.5% chlorine solution mixed at 08:00 is essentially weak salt water by 10:00. The fix: mix fresh every two hours, or work from concentrate and dilute at the point of use. Most teams skip this: they fill sprayers in the morning and assume the solution holds all day. It doesn't. The trade-off is labor—mixing small batches repeatedly eats time—but the alternative is spraying colored water onto contaminated surfaces. That hurts.

Store concentrate in opaque containers. Light accelerates decay.

Do I need to rinse after using a disinfectant?

It depends on the surface and the disinfectant class. For chlorine on non-porous surfaces in a cholera treatment center: no rinse—let it air-dry. Chlorine residue continues working until it evaporates. Rinsing with water removes the active chemical and re-wets the surface, potentially spreading pathogens. However—and this is where the protocol gets muddy—if the surface touches food or a patient's mouth, rinse with potable water after the required contact time. Bleach residue can irritate mucous membranes and corrode metal instruments over weeks of repeated application.

Quat-based disinfectants often leave a sticky film that traps dirt. Some national guidelines mandate rinsing after 10 minutes. Others say leave it. Check your local protocol before you assume. What usually breaks first is staff time: nobody wants to wait ten minutes, then wipe, then rinse. So they skip steps. The better answer is choosing a disinfectant that matches your workflow, not the other way around.

'We stopped rinsing because we ran out of clean water. The infection rate didn't go up. But the zinc buckets rusted through in three months.'

— WASH officer, South Sudan field rotation, 2023