Eco‐Microbiology: A Frugal‐Circular Framework for Biosafe, Low‐Cost Practical Microbiology in Secondary Education

ABSTRACT

Microbiology offers valuable experiential learning, but secondary‐school practice is limited by cost, infrastructure, biosafety and teacher confidence. This review presents a framework for sustainable, frugal‐circular and pedagogically rigorous microbiology in resource‐limited settings, positioning frugality not only as low‐cost provision but as a teachable behavioural principle of minimal resource extraction, maximal conservation and upstream waste prevention. It shows how to replace standard equipment and consumables with accessible, reusable alternatives (e.g., domestic pressure cookers, improvised incubators, everyday glassware) while maintaining equity, scientific rigour and clear biosafety boundaries. Circular science education is operationalised through guidance on constructing essential tools, preparing culture media from household ingredients and choosing safer, food‐associated microbial sources (fermented foods, probiotics) appropriate for biosafety level 1 teaching. When environmental materials are used, activities are limited to teacher‐led, sealed observation and the precautionary rule ‘unknown = potentially pathogenic’. The paper provides protocols for sterilisation, disinfection and waste management, including an end‐of‐session workflow for reliable culture inactivation before disposal. An implementation guide and checklist translate these principles into classroom routines, with a model experiment, assessment ideas and management strategies. The framework demonstrates that inclusive, environmentally responsible microbiology is feasible across diverse school contexts and can strengthen scientific citizenship by linking laboratory practice to global stewardship concepts, including frugality.

Eco‐Microbiology: a frugal‐circular pedagogical framework for biosafe, low‐cost, low‐impact microbiology in secondary education—This graphical abstract summarises the Eco‐Microbiology framework, which integrates frugality as a behavioural and design principle (minimal extraction, maximal conservation and upstream waste prevention) alongside biosafety, instructional rigour and circular resource use in school microbiology.

1. Introduction

Microbiology is a fundamental branch of science that enables students to engage directly with the living microbial world. Practical laboratory experiences are particularly effective in helping learners develop scientific reasoning, investigative skills and an awareness of microbiological processes that influence health, food and the environment (e.g., Gericke et al. 2022; Apeadido et al. 2024; Kumari et al. 2024; Timmis et al. 2024). However, secondary schools frequently encounter substantial barriers to implementing hands‐on microbiology teaching, notably due to the cost, limited availability and environmental footprint of standard laboratory materials and equipment (e.g., Chala 2019; Amorim, Lopes, et al. 2025).

In many educational contexts (especially in under‐resourced or rural schools), despite teacher and pupil enthusiasm and thirst for hands‐on experimental exercises, access to professional laboratory infrastructure remains limited (e.g., Drescher et al. 2022; Khethiwe 2023; Shambare and Jita 2025). In parallel, the growing imperative to align education with ecological responsibility calls for rethinking the widespread reliance on disposable plastics and resource‐intensive tools commonly used in microbiological procedures (e.g., Alves et al. 2021; Yusuf et al. 2022; Carrillo‐Barragan 2024; Bayrau et al. 2025). Collectively, these constraints often lead to microbiology being delivered primarily at a theoretical level, depriving students of essential experiential learning opportunities and reinforcing inequities in science education (e.g., Sshana and Abulibdeh 2020; Smith‐Keiling 2021).

Despite increasing attention to sustainable laboratory practices, a comprehensive pedagogical framework for integrating low‐cost, accessible and ecologically responsible microbiology into secondary education is still lacking. This article addresses that gap by proposing an integrated framework for safe, ecologically responsible and low‐cost microbiological experimentation in secondary schools. Drawing on practical adaptations using household or readily available materials, we present sustainable alternatives to conventional laboratory tools and consumables that preserve educational value and maintain biosafety. These strategies aim to foster a culture of reuse, creativity and environmental stewardship, while also developing awareness of profligate practices and avoidable waste and expanding equitable access to promote teacher agency towards meaningful microbiology learning.

More broadly, school curricula may be highly compartmentalised for understandable practical reasons, and cross‐cutting issues such as biosphere stewardship, stakeholder responsibilities, conservation and sustainability may therefore be under‐emphasised or omitted because an appropriate subject context is not always apparent. Microbiology, however, offers a multitude of authentic contexts to make these issues explicit—linking everyday practices (resources, waste, risk) to responsible decision‐making.

By linking science teaching with sustainable practice, this work supports broader educational and environmental objectives, including the United Nations Sustainable Development Goals (SDGs), notably SDG 4 (Quality Education) and SDG 12 (Responsible Consumption and Production) (United Nations 2015). Here, ‘frugality’ is used as a pedagogical and behavioural principle—careful, minimal resource use and upstream waste prevention—rather than as a synonym for ‘cheap’, consistent with work on frugal education design (Masters 2024).

1.1. Transferability Note

Examples of ‘household’, ‘food‐grade’ or ‘retail‐available’ materials in this paper are illustrative rather than prescriptive. Teachers should substitute locally available equivalents (e.g., local markets, cooperatives, pharmacies, school suppliers) and document product type/grade and any relevant batch variation when reproducibility matters. This flexibility can itself be treated as a pedagogical opportunity: teachers and students may critically evaluate and refine alternative materials, using the decision criteria outlined in Section 3.3 to assess biosafety, rigour and suitability.

1.2. Optional Field‐Facing Extension

Microbiology is also frequently conducted in field settings or begins with field observation and sampling. Class excursions can therefore provide motivating, place‐based opportunities for observation‐first activities that connect learners with microbial habitats and biodiversity, with documented educational value (McGenity et al. 2020; Archer et al. 2025). Field work is best framed primarily as observation and simple environmental measurement (e.g., temperature or pH gradients), including portable or home‐made microscopy approaches where appropriate (IMiLI_Mic). Where environmental samples are collected for follow‐up work, they should be treated within the same precautionary boundary used throughout this framework (‘unknown = potentially pathogenic’) with any subsequent culturing restricted to teacher‐led, sealed handling and validated end‐of‐session inactivation.

The following sections provide practical equipment substitutions, alternative culture‐media options, low‐risk microbial sources and implementation strategies designed to support teachers' agency in integrating microbiology education with curricular goals across diverse classroom environments.

Ultimately, this paper offers a novel, integrative approach to microbiology teaching that combines hands‐on pedagogy with ecological and ethical responsibility. Rather than offering isolated low‐cost hacks, we articulate a decision framework that specifies when frugal substitutions are acceptable, what constitutes sufficient methodological rigour in secondary school settings, and which biosafety boundaries are non‐negotiable. In particular, we make explicit a set of minimum measurement requirements appropriate to this level: basic control and recording of incubation parameters (time and temperature), estimation or measurement of pH where relevant, the systematic use of replication and negative controls, and structured documentation of procedures and deviations. These criteria ensure that classroom observations remain interpretable even when equipment is simplified. Beyond technical robustness, they also model core elements of the scientific method and ethos—rigour in planning and execution, systematic control of variables, agnostic interpretation of results, transparency in documentation and explicit recognition of uncertainty. Operationally, these criteria are implemented through the risk/decision matrix (Table S4), which functions as a biosafety and feasibility gatekeeper, guiding activity selection, permitted manipulations and end‐of‐session inactivation and waste management.

2. Challenges in Implementing Practical Microbiology Work Using Sustainable Tools in Secondary Education

Despite the recognised pedagogical value of hands‐on laboratory work in microbiology (e.g., Fahnert 2017; Coyte 2023; Moreno et al. 2023; Prate and Hsu 2025), secondary school settings often encounter significant barriers to its effective implementation (e.g., Spell et al. 2014; Petersen and Chan 2020; Gericke et al. 2022). These challenges span logistical, financial, pedagogical and safety‐related dimensions, which collectively constrain teaching effectiveness and student engagement.

One of the most immediate and pervasive barriers is the lack of appropriate, safe and affordable resources, including: (Abedi‐Firoozjah et al. 2022) laboratory facilities, equipment and consumables; (Aliyo and Edin 2023) access to low‐risk microorganism strains; and (Alraddadi et al. 2023) standard culture media or clear guidance on how to prepare them. Many secondary schools worldwide lack essential microbiological tools, such as incubators, autoclaves, high‐quality microscopes or sterile workspaces. Even when basic laboratory settings exist, limited access to consumables—such as Petri dishes, agar media, sterile pipettes, gloves and disinfectants—compromises the reproducibility and reliability of experimental outcomes and restricts the routine implementation of laboratory practices with students. Inadequate facilities, including poorly ventilated rooms and insufficient workspace, further constrain the scope of microbiological investigations that can be carried out safely and effectively (e.g., Pareek 2019; Bouzit et al. 2023; Lloyd et al. 2023; Van Dat et al. 2023; Mangarin and Macayana 2024).

Time constraints imposed by standard class schedules (e.g., Teig et al. 2019; Batty and Reilly 2022) also limit the feasibility of experiments that require incubation or sequential observation over multiple days. In addition, the appropriate disposal of biological waste poses logistical and regulatory challenges, particularly when working with live cultures of environmental microorganisms whose pathogenic potential cannot be excluded. In many schools, biosafety protocols are underdeveloped or inconsistently enforced, raising concerns about student safety awareness and environmental contamination (e.g., Smith‐Keiling 2021; Aliyo and Edin 2023; Mendez 2023). More broadly, the perceived risk associated with handling live microorganisms—whether driven by health concerns or institutional liability—can lead to overly cautious or entirely theoretical approaches to microbiology education. This challenge is compounded by a limited biosafety culture: students may be unfamiliar with basic hygiene practices or containment strategies, increasing the risk of cross‐contamination or accidental exposure (e.g., McMichael 2019; Nwachukwu and Nnagbo 2024; Rezal et al. 2024).

From a pedagogical perspective, teachers often lack specific and updated training in microbiological techniques, making it difficult to guide students confidently through complex procedures or to troubleshoot technical problems (e.g., Fidiastuti et al. 2024; Valderrama et al. 2025). Likewise, students often require explicit scaffolding to follow experimental protocols reliably, maintain basic biosafety/aseptic routines, and record and interpret observations as evidence (e.g., Maicas et al. 2020; Chengere et al. 2025). There is frequently a disconnect between theoretical content and laboratory practice, which can reduce the perceived relevance and educational impact of experiments. Furthermore, limited exposure to scientific methodology may contribute to a superficial understanding of experimental design and data analysis (Kranz et al. 2023). Yet it is precisely engagement with the scientific process—formulating testable questions, using controls, recognising uncertainty and interpreting evidence—that provides transferable lessons central to scientific literacy and citizenship.

Beyond the laboratory itself, institutional and systemic factors also play a role. These include insufficient administrative support (Lotter et al. 2024), inadequate funding for science education (Fitzgerald et al. 2019; Cayton and Jones 2023) and bureaucratic obstacles related to the acquisition and handling of biological materials. Strict legal regulations governing the use of microbial agents in educational settings—although essential for ensuring safety—may nevertheless discourage teachers from implementing low‐risk practical instructional approaches (Byrd et al. 2019; European Union 2000), thereby limiting exploratory learning and reducing opportunities for inquiry‐based pedagogy.

Finally, students tend to be more deeply engaged when given autonomy over experimental design and inquiry, whereas prescriptive or ‘cookbook’ laboratory activities can reduce engagement due to a lack of ownership or perceived relevance (Marley et al. 2022; Yang et al. 2022; Cirkony et al. 2025). When practical microbiology is reduced to traditional, rote‐based laboratory methods, learners may have fewer opportunities to develop science process skills and critical thinking, which can undermine engagement; conversely, explicitly foregrounding societal relevance is key to awakening curiosity and helping students recognise real‐world applicability (Chengere et al. 2025; Timmis 2023b). Here, we examine solutions for implementing a low‐cost school laboratory, with particular emphasis on environmentally sustainable options.

3. Pedagogical Framing: Eco‐Microbiology and ‘Circular Science Education’

Microbiology is well suited to sustainability‐oriented science education because it links cellular processes to food systems, environmental cycles, hygiene and public health. In secondary classrooms, an eco‐microbiology framing can therefore treat practical work not only as a technical exercise, but also as a context for reflecting on resource use, waste, risk management and responsibility in scientific practice. This orientation aligns with sustainability‐focused educational scholarship that calls for learning designs that are resource‐conscious, adaptable to local constraints and explicitly connected to wider societal challenges (e.g., Hays and Reinders 2020; Gamage et al. 2022; Ramos et al. 2023; Tiippana‐Usvasalo et al. 2023; Timmis et al. 2024; Amorim, Santos, and Timmis 2025).

In education design, frugality is increasingly framed as a proactive mode of leveraging available resources (time, space, materials) while reducing unnecessary consumption, rather than a reactive response to scarcity. This framing supports positioning microbiology practical activities as an explicit site for teaching resource stewardship and responsible practice (Masters 2024).

3.1. Guiding Principles: Rigour, Equity, Sustainability, Biosafety

• Scientific rigour|Low‐cost approaches should not imply lower methodological standards. Rigour can be maintained through explicit protocols, careful labelling, routine use of controls (e.g., uninoculated media; ‘air‐exposed’ plates) and structured observation and recording. Where feasible, simple replication and comparison of conditions can be used to strengthen inference and improve students' understanding of evidence‐based claims.

• Educational equity|Practical microbiology can amplify inequities when access to equipment, consumables and safe cultures is uneven across schools. A key aim of circular approaches is therefore to broaden access to authentic laboratory experience by proposing substitutions and procedures that remain pedagogically robust while being feasible in resource‐limited contexts.

• Teacher agency (situated judgement)|Teachers act as real‐time risk and quality managers, making situated decisions (e.g., deviations, containment failures, unexpected growth) and turning these moments into explicit learning about uncertainty, controls and responsible practice.

• Environmental sustainability|Conventional microbiology practical classes often depend on single‐use plastics and resource‐intensive consumables. Circular approaches prioritise reuse, repurposing and low‐impact methods (where compatible with safety), encouraging students to examine trade‐offs and the environmental footprint of laboratory practice, and to appreciate important sustainability principles.

• Biosafety and responsibility|Biosafety is non‐negotiable in any microbiology activity. Frugal‐circular classroom adaptations must therefore operate within clear biosafety boundaries, including organism choice, containment, disinfection/sterilisation and waste management, so that curiosity is paired with risk awareness and reliable end‐of‐session inactivation routines.

3.2. What ‘Circularity’ Means in School Science: Reuse, Low Impact, Local Adaptation and Curriculum Alignment

In a school laboratory context, circularity can be defined operationally as: designing practical activities that minimise waste and resource consumption while maintaining educational value and biosafety. This definition emphasises three practical elements: (i) reuse and repurposing of materials that can be cleaned and sterilised safely; (ii) low environmental impact, including reduction of single‐use plastics and unnecessary hazardous chemicals; and (iii) local adaptation, whereby activities are designed around locally available resources and culturally familiar microbial sources (e.g., fermented foods), reducing dependence on specialised supply chains (Nielsen 2017; Hays and Reinders 2020; Tiippana‐Usvasalo et al. 2023). Importantly, this framework is intended to support—not compete with—curriculum delivery: practical microbiology is used as a vehicle to teach required learning outcomes and cross‐curricular competences (e.g., experimental design, measurement, data interpretation and health/environment connections), while adapting materials and routines to local constraints within non‐negotiable biosafety boundaries.

At the secondary level, this framing also supports explicit teaching about why circular practices require deliberate criteria and routines rather than ad hoc improvisation—for example, specifying what can be reused, how it is cleaned or sterilised, and how uncertainty is handled when working with mixed or variable microbial sources. Education research on circular economy learning in schools highlights both the relevance of these ideas and the need for structured didactic approaches appropriate to student levels (e.g., Keramitsoglou et al. 2023; Laius et al. 2024). In this context, circularity in school microbiology cannot be reduced to improvised substitutions; it requires explicit pedagogical structure, clearly articulated biosafety boundaries and transparent criteria for responsible adaptation.

3.3. Decision Criteria for Acceptable Substitutions

Translating this framing into practice requires explicit decision criteria to guide the adoption of alternative methods, materials and devices before classroom implementation. A substitute approach should be used only if it satisfies four conditions:

1. Biosafety boundary|it must not increase exposure risk (spills, aerosols, open handling, uncontrolled incubation) and must remain compatible with the activity's risk classification, decontamination route and waste handling. Here, ‘exposure risk’ refers to plausible routes of contact or entry (e.g., hand‐to‐mouth/ingestion, inhalation of aerosols, splashes to skin/eyes) and is mitigated through behavioural controls and containment routines (see Section 8 and Table S4).

2. Learning‐objective integrity|it must preserve the core conceptual phenomenon (i.e., the intended learning goal) targeted by the lesson so that results remain interpretable within the intended conceptual scope.

3. Fit‐for‐purpose evidence|it must support the intended strength of inference (e.g., descriptive/qualitative vs. comparative/semi‐quantitative), with appropriate controls and without implying precision the method cannot deliver, a transferable lesson in evidence‐based reasoning that extends beyond microbiology.

4. Classroom reproducibility|it must enable within‐class replication and minimal documentation of key inputs and conditions (e.g., source/material, container type, time, temperature and any deviations) sufficient to interpret variability responsibly (see Section 9.6)

These criteria function as a gatekeeping tool for local adaptation; remaining constraints and performance limits are discussed separately in the Limitations section. These criteria are also implemented in practice through the stepwise checklist in the Supporting Information (Table S4), which standardises activity selection, permitted manipulations, decontamination routes and waste handling procedures. A concise set of baseline Good Microbiological Practice (GMP) measures applies across all activities, irrespective of specific substitutions. Baseline Good Microbiological Practice (GMP) measures include:

• –Use of protective clothing (lab coat or dedicated apron), closed footwear and tied‐back hair.
• –No eating or drinking in the workspace.
• –Strict prohibition of placing any objects (including writing instruments or fingers) in the mouth while in the laboratory, recognising ingestion as a primary exposure route.
• –Clear separation between experimental and non‐experimental materials.
• –Surface disinfection before and after activities.
• –Proper hand hygiene before and after the session.
• –Sealed incubation and prohibition of reopening cultures.
• –Explicit end‐of‐session confirmation of containment, waste segregation and workspace cleaning.

3.4. Didactic Implications: Interdisciplinarity, Student Agency, Scientific Citizenship

• Interdisciplinarity and systems thinking|Eco‐microbiology naturally integrates biology with chemistry (pH, indicators, metabolism), physics (temperature control, optics/microscopy), and environmental science (waste, contamination pathways, life cycle thinking), while also connecting to applied domains such as food science (fermentation and safe cultures), health sciences (hygiene practices, biosafety procedures, link to public health) and climate‐related processes (e.g., microbial contributions to CO₂ or CH₄ production). Making these connections explicit supports systems‐oriented understanding and helps students relate laboratory decisions to broader ecological consequences.

• Student engagement and inquiry|Practical microbiology work lends itself to inquiry‐based learning because students can test, compare and refine protocols and conditions (e.g., comparing media formulations, evaluating indicator performance, assessing temperature effects, investigating optimal time of results readout or comparing microbial sources). Frugal‐circular adaptations can further support this inquiry by enabling structured comparisons of alternative materials, devices and workflows, while keeping learning objectives and biosafety boundaries explicit. Framing constraints and variability as design parameters can support problem‐solving and a sense of ownership over the investigation within teacher‐scaffolded boundaries, strengthen understanding of controls and reproducibility, and help learners appreciate why outcomes in biological systems can vary even when procedures are standardised. A useful pedagogical distinction can be made between discovery‐oriented observation (pattern noticing) and question‐driven experimentation; the former often motivates inquiry, while the latter frames purpose and interpretation without requiring additional technical complexity. In secondary‐school microbiology, question‐driven experiments can be built around a small set of high‐leverage environmental variables that are conceptually meaningful and feasible to control in resource‐constrained settings—for example pH (selection pressure and niche constraints), temperature (enzyme activity and growth kinetics), oxygen availability (aerobic vs. microaerophilic growth) and substrate composition (nutrient availability and selective bias). Keeping other parameters constant (e.g., inoculum dilution, incubation time and container type) supports interpretive rigour while preserving a low‐barrier design.

• Scientific citizenship|Microbiology education is increasingly framed as societally relevant, supporting informed decision‐making about health, environment and technology across educational levels. Recent work has extended microbiology education into early childhood settings, demonstrating that carefully designed, inquiry‐oriented experiences can foster curiosity and foundational microbial literacy even before formal secondary schooling (Di Capua et al. 2026). This orientation aligns with microbiology literacy initiatives that emphasise the civic importance of understanding microbes and microbial processes (e.g., Anand et al. 2023; Timmis 2023a; Timmis et al. 2024; Amorim, Santos, and Timmis 2025). In parallel, research on citizen science in education indicates that carefully scaffolded participation can support learning outcomes and civic engagement, particularly when linked to authentic, contextualised investigations (e.g., Finger et al. 2023; Hadjichambi et al. 2023; Solé et al. 2024).

Beyond civic literacy, the procedural habits cultivated through frugal‐circular microbiology—such as documentation discipline, quality control reasoning, resource‐aware decision‐making and explicit management of variability—mirror foundational competences required in biotechnology and emerging bioeconomy sectors. Even simplified classroom investigations require scheduling of incubation periods, coordination of parallel steps and allocation of effort across sessions, making visible the organisational dimension of scientific practice. While the framework does not simulate industrial production, it introduces students to structured thinking about biological processes, constraints and standardisation that underpin responsible biomanufacturing contexts.

Selected transferable ‘life lessons’ that can be explicitly taught through microbiology practical activities are summarised in Table S5.

4. Practical Solutions I—Essential Equipment: Sustainable and Low‐Cost Alternatives

Implementing microbiological practical work in secondary education is often constrained by the high cost, maintenance demands and environmental impact associated with conventional laboratory equipment. Although several low‐cost solutions are already available, many do not sufficiently address environmental sustainability. There is therefore a need for a comprehensive range of low‐cost and sustainable alternatives that can be adopted without compromising biosafety or pedagogical value. Embedding sustainability considerations into alternative laboratory practices can help students develop, from an early age: (i) awareness, values and attitudes aligned with sustainability; and (ii) creativity and problem‐solving skills through the application of scientific principles using inexpensive, environmentally responsible materials—especially when the teacher explicitly compares conventional laboratory practices with frugal‐circular substitutions and discusses their resource and waste implications (see Table S5).

This section outlines feasible substitutions for eight essential microbiological tools: the autoclave, incubator, flame source, aseptic work area, microscope, anaerobic jar, centrifuge and pH metre. Readers may also complement the guidance below with reputable online resources, including instructional videos and images. For each tool below, we focus on function, a validated school‐appropriate substitute, minimum essential performance checks and safety constraints.

4.1. Sterilisation: Autoclave → Domestic Pressure Cooker

Autoclaves are widely used to sterilise culture media, glassware and instruments; however, their cost and operational requirements make them inaccessible to many secondary schools. A domestic pressure cooker can provide a practical alternative. When operated under appropriate conditions, pressure cookers (Figure 1a) can reach internal temperatures of approximately 121°C at around 15 psi, which is generally sufficient to inactivate microbial contaminants, including bacterial endospores (Swenson et al. 2018). To ensure safe and effective use in school settings, teachers should provide explicit instruction and supervision, emphasising basic safety precautions (e.g., ensuring the lid is correctly locked before heating; reducing heat once operating pressure is reached; and allowing pressure to fall naturally to atmospheric pressure before opening the valve). When bags are used for sterilisation or decontamination, they must be made of heat‐resistant material suitable for pressure/steam exposure and should not be assumed to be interchangeable with ordinary plastic bags. Likewise, glass bottles or jars containing liquids should not be filled beyond approximately three‐quarters of their total volume and should not be tightly sealed during the cycle, in order to reduce pressure‐related risks. Where available, institutional guidance for school laboratory practice can support safe implementation (e.g., CLEAPSS guidance on the use of pressure cookers in school science: cleapss.pressure‐cooker).

FIGURE 1.

Low‐cost, biosafe substitutes for core microbiology equipment in secondary‐school settings. (a) Domestic pressure cooker on a hotplate used for steam sterilisation; (b) improvised incubator: Insulated box fitted with a low‐wattage heat source; (c) improvised aseptic work box (still‐air box) using a homemade spirit lamp and a metal‐wire inoculation loop; (d) candle‐jar set‐up to generate a reduced‐oxygen/CO₂‐enriched atmosphere; (e) cotton plugs used as reusable closures for culture tubes; (f) liquid‐transfer substitutes—needleless syringes and glass droppers—used alongside reusable household gloves. Do not use disposable plastic gloves near open flames; prefer hand hygiene for low‐risk BSL‐1 activities. Refilling is teacher‐only, with the flame extinguished and away from ignition sources.

4.2. Incubation: Incubator → Insulated Box With Heat Source

Maintaining microbial cultures at appropriate incubation temperatures (typically 30°C–37°C) is fundamental to many microbiology activities (Madigan et al. 2021). Where a dedicated incubator is unavailable, an insulated container (e.g., an insulated foam or plastic cooler) can be paired with a low‐wattage heat source such as an incandescent light bulb (15–40 W) or a rechargeable heating pad (Figure 1b). When connected to a simple thermostat or digital temperature controller (e.g., an Inkbird‐type controller), such systems can provide stable, adjustable heating that approximates the function of a laboratory incubator. Where possible, the set‐up should be tuned to maintain temperature stability within approximately ±1°C–2°C and verified using an independent thermometer before use with cultures. To minimise fire and burn risks, any materials in direct contact with, or near, the bulb must be heat‐resistant and the bulb should be positioned with adequate clearance and shielding to prevent contact with plastic surfaces or paper‐based insulation. Where available, institutional or publicly available guidance for classroom engineering controls can support safe construction and testing (e.g., open educational engineering resources on insulated incubator boxes: teach engineering‐insulation).

Low‐cost and open‐source incubation solutions have also been validated in peer‐reviewed studies. For instance, Arumugam et al. (2021) described a portable, low‐cost CO₂ incubator built using 3D‐printer hardware; Samokhin et al. (2022) demonstrated an inexpensive system capable of regulating temperature, O₂ and CO₂ for cell culture; and Wight et al. (2020) developed a rugged, cost‐effective field incubator for microbiological analyses that performed comparably to commercial units. These studies support the feasibility of simplified incubation systems for teaching contexts, provided that performance is verified independently and documented for each session.

4.3. Flame Source: Bunsen Burner → Alcohol Lamp (Spirit Lamp)

An open flame is commonly used to support aseptic technique, including the flame sterilisation of inoculating loops, briefly flaming the mouths of vessels and helping to maintain a low contamination working area. In settings where gas infrastructure is unavailable, restricted or presents additional safety risks, alcohol (spirit) lamps can provide a practical substitute for a traditional Bunsen burner (Figure 1c). These refillable devices—typically fuelled with ethanol or isopropanol—produce a stable flame that can be readily controlled for routine aseptic handling and the flaming of small instruments.

Disposable plastic gloves should not be used when working with open flames due to flammability and burn risk. In low‐risk Biosafety level 1 (BSL‐1) teaching activities, hand hygiene before and after the session is generally preferable to glove use. Where gloves are required by local policy, PPE selection and workflow should minimise ignition hazards, and fuel handling/refilling must be teacher‐only and performed with the flame extinguished and away from ignition sources.

The use of alcohol lamps as a heat source is well documented across microbiological and biomedical contexts. For example, Kempraj et al. (2024) describe flaming forceps using a spirit lamp during aseptic dissections; Zhang et al. (2025) report protocols in which an alcohol lamp is used to sterilise instruments between handling steps; and Huang et al. (2025) describe fixing slides by briefly passing them through an alcohol‐lamp flame. Collectively, these reports indicate that spirit lamps are a widely used alternative to Bunsen burners for routine aseptic manipulations.

From a classroom perspective, the use of spirit lamps also provides a structured opportunity to teach and reinforce laboratory safety. Teachers should ensure active supervision and explicit instruction on safe handling of flammable liquids (e.g., using small volumes; keeping fuel containers closed and away from the flame; tying back hair and securing loose clothing; maintaining a clear, uncluttered bench; and extinguishing the lamp by replacing the cap rather than blowing it out).

4.4. Aseptic Area: Laminar Flow Hood → Improvised Aseptic Work Box

Although laminar flow cabinets are standard in professional laboratories, their cost and technical requirements are often prohibitive in secondary education settings. A practical alternative is a still‐air box or glove box, constructed from a transparent, sealable container fitted with glove ports to enable manipulation within a semi‐controlled environment. While this arrangement does not provide HEPA filtration, it can create a stable, low‐draught workspace that reduces airborne contamination during procedures such as inoculation and plating. Studies have reported the utility of glovebox and still‐air box designs for semi‐sterile operations and controlled‐atmosphere applications (e.g., Kosel and Ropret 2021; Pereira et al. 2023).

Simpler enclosures, such as a medium‐to‐large aquarium or a wooden box oriented with the opening towards the operator, may also support cleaner handling, provided that good aseptic technique is maintained (Figure 1c vs. Figure 1f).

An ultraviolet (UV‐C) lamp can be used to reduce airborne and surface contamination within a closed aseptic enclosure prior to a microbiology class, particularly in resource‐limited settings where laminar flow systems are unavailable. Where UV‐C irradiation is used, it must be operated within a fully closed enclosure and strictly in the absence of students and other personnel. UV‐C devices should be used exclusively for short pre‐disinfection periods (e.g., 15 min) and must be switched off before any manipulation begins. Because UV‐C radiation poses significant eye and skin hazards—and some lamps may generate ozone—human exposure must always be prevented, (e.g., through the use of a closed enclosure, timer or interlock where available), and appropriate ventilation must be ensured where relevant. In routine use, enclosure surfaces should also be cleaned with an alcohol‐based spray before and after each session. Together, these measures provide a simple and reusable approach to supporting aseptic conditions in teaching and in other resource‐limited laboratory contexts.

4.5. Microscopy: Portable/Open‐Source Solutions and Online Resources

Optical microscopes remain central to microbiology education. In secondary school contexts, use of laboratory‐grade instruments may be limited by infrastructure, maintenance or procurement constraints. Nevertheless, functional entry‐level microscopes are increasingly available at substantially lower cost through widely accessible retail channels, and several online platforms provide interactive microscopy resources that teachers can use at a level of complexity appropriate to their students (e.g., microscopyu; evidentscientific; virtuallabs).

In parallel, advances in open‐source and portable optical systems have made high‐quality microscopy more accessible. For example, the OpenFlexure microscope provides a 3D‐printed, low‐cost platform for bright‐field and fluorescence imaging and can be assembled using recycled or locally sourced components (Collins et al. 2020). Similarly, recent developments in mobile and modular optical microscopy—including smartphone‐integrated systems and adaptable lens modules—enable quantitative imaging of microorganisms at a fraction of the cost of traditional instruments (Liu et al. 2021; Ganesan et al. 2022). Ultra‐low‐cost educational designs such as foldscopes and structured home‐made microscopy approaches (e.g., IMiLI_Mic), further extend accessibility by enabling direct observation through paper‐based or improvised optical assemblies. These approaches are particularly valuable in field‐based, outreach and resource‐limited teaching contexts and align with broader goals of inclusive and sustainable science education. Such designs are typically lightweight, repairable and energy‐efficient and they lend themselves to interdisciplinary school projects that integrate biology with physics and engineering, while also supporting practical microbiology education and citizen science.

4.6. Anaerobic/Microaerophilic Atmosphere: Sealed Jar With Candle Method

Cultivating microorganisms under anaerobic or microaerophilic conditions typically requires a dedicated anaerobic jar with gas‐generating systems to reduce oxygen availability. As a practical alternative for school‐based demonstrations, a hermetically sealed glass or metal container used with a small candle can generate a low‐oxygen, CO₂‐enriched atmosphere suitable for basic educational purposes (Figure 1d). In this approach, inoculated culture plates are placed inside the container alongside a lit tea light or small candle. Once sealed, the flame consumes oxygen and extinguishes when the oxygen concentration falls below the level required to sustain combustion, thereby reducing oxygen availability and increasing carbon dioxide. This atmosphere can support the growth of certain facultative anaerobes and microaerophiles, and it can be particularly useful for illustrating the effects of oxygen limitation in microbial growth.

Several BSL‐1 organisms commonly associated with food fermentations respond well to reduced‐oxygen, CO₂‐rich conditions. For example, Lactobacillus spp. (found in yoghurt, kefir and fermented vegetables), Streptococcus thermophilus (used in dairy fermentations) and Leuconostoc spp. (associated with kimchi and sauerkraut) can grow well in low‐oxygen environments (Carr et al. 2002; Shaikh et al. 2021). These microorganisms are therefore suitable for educational demonstrations using simple, accessible materials, provided that the activity is conducted under close supervision and with appropriate precautions (e.g., using a heat‐resistant container, keeping flammable materials clear and ensuring the flame is fully extinguished before opening).

4.7. Centrifugation (Optional): Manual Alternatives

A centrifuge is basic equipment in biology laboratories and is commonly used for sedimentation and short spin‐down steps. In school contexts, access may be limited to simpler or improvised systems rather than high‐speed laboratory instruments. For educational purposes, where high g‐forces are not required, low‐speed manual alternatives can be assembled from repurposed kitchen devices—most commonly a salad spinner adapted to hold tubes, racks or plates—enabling basic sedimentation or short spin‐down steps. Such approaches have been described as practical teaching microcentrifuges (Moran and Galindo 2011), explored as hand‐powered centrifugation in applied contexts (Brown et al. 2011), and more recently demonstrated for brief spin‐downs of solutions in 96‐well PCR plates using a commercially available salad spinner (Motohashi 2020).

4.8. pH Estimation, Adjustment and Monitoring: Handheld Metres, Indicator Strips and Built‐In Medium Indicators

Microbial growth is constrained by pH, and many school investigations benefit from either adjusting media prior to use or monitoring pH shifts driven by microbial metabolism. Handheld pH metres can provide sufficiently rigorous readings for classroom work when treated as quantitative instruments. At minimum, metres should be calibrated at the start of each practical session using at least two buffers that bracket the expected pH range, with careful electrode rinsing/blotting between buffers and samples (see ‘minimum QC for handheld pH measurement’ in Supporting Information document Box S1 for details). When calibration buffers are not available, values should be reported explicitly as estimates, interpreted primarily through relative comparisons, and (where possible) cross‐checked with indicator strips.

Where visual monitoring is desirable, indicator systems can be incorporated directly into the culture medium to make otherwise abstract changes observable. For example, anthocyanin‐rich red cabbage extract has been reported in indicator‐incorporated media as a pH‐responsive colourimetric system (Celik et al. 2020). In more traditional microbiology teaching, litmus milk similarly embeds an indicator system within a nutrient medium, enabling learners to infer metabolic activity through characteristic colour changes (e.g., Qian et al. 2018; Salehzadeh and Kheirjou 2024). Phenol red is widely used as a built‐in pH indicator in carbohydrate fermentation media, and other indicators (e.g., bromothymol blue and bromocresol purple) can be selected to match the expected pH transition range and instructional aim (see, for example, microbenotes). In addition, some demonstrations may incorporate redox indicators such as methylene blue, which changes colour as oxygen is depleted and reducing conditions develop—thereby complementing pH‐based observations by illustrating how microbial activity can simultaneously alter the chemical environment through both acid–base and oxidation–reduction processes (e.g., Roy et al. 2024). Practical guidance for preparing and incorporating natural indicators (including plant‐based options) is provided in the Section 7 Low‐cost culture media and natural indicators, to avoid duplication and to keep pH estimation procedures distinct from media formulation.

5. Practical Solutions II—Consumables and Materials: Reusable Substitutions

In addition to equipment, microbiology education at secondary level depends heavily on consumable materials, many of which are single‐use, plastic‐based or costly. To support both ecological responsibility and accessibility in school laboratories, it is essential to identify practical, low‐cost alternatives that can be reused, sterilised and adapted from everyday materials. Below, we outline several such substitutions, demonstrating how essential microbiological procedures can be maintained through more sustainable practices (Freese et al. 2024; Roy and Chakraborty 2024). A complementary strategy is to establish with nearby organisations that operate microbiology or analytical laboratories. In addition to universities and research institutes, relevant partners may include local or national industries, government laboratories (e.g., food safety or environmental testing facilities), hospital diagnostic laboratories and other public‐sector laboratories. Such partnerships can facilitate the responsible transfer or repurposing of surplus or decommissioned laboratory materials (e.g., glass Petri dishes, pipettes, test tubes and small equipment) that remain suitable for safe educational use in schools. Beyond material support, these collaborations can foster outreach, strengthen school–laboratory connections and expose students to real‐world applications of microbiology.

5.1. Petri Dishes → Reusable Glass Jars/Lids

Petri dishes are essential for culturing microorganisms, ‘dilution‐to‐extinction’ and distinguishing different microbes in samples are typically made of glass or plastic. Although glass Petri dishes are more expensive, they are reusable and sterilizable and therefore generally more sustainable than disposable plastic dishes. Plastic Petri dishes are widely used in research laboratories because they can be purchased pre‐sterilised in bulk; however, for schools, glass Petri dishes may represent a more suitable long‐term option.

As an alternative, small glass jars—such as those used for yoghurt, jam or spices—can be repurposed as culture containers. Many are sufficiently heat‐resistant and can be readily sterilised (e.g., using a pressure cooker), while their transparency allows microbial growth to be monitored. In some cases, the metal or glass lids can serve as shallow culture platforms, covered with the inverted jar to create a simple humid chamber that helps reduce desiccation and contamination. These approaches can minimise plastic waste and substantially reduce costs (Trusler et al. 2024). As a precaution, jars and lids should be inspected for cracks, chips or degraded seals and only items known to tolerate heat exposure should be used for sterilisation. In addition, low‐cost and environmentally preferable covers can be prepared using cotton wool wrapped in one or two layers of gauze, with the size adjusted to fit the diameter of the flask or tube.

5.2. Inoculating Loops → Metal Wire/Shaped Paperclips

An inoculating loop is an essential tool for aseptic technique, yet it can be readily replaced with a handmade loop formed from stainless‐steel wire, straightened paper clips or copper wire, shaped and mounted on a reusable handle (e.g., a cork or wooden stick) (Figure 1c). The metal loop can be flame‐sterilised and reused repeatedly, providing a durable, low‐cost option that substantially reduces single‐use waste (Alves et al. 2021; Carrillo‐Barragan 2024). As a precaution, any cut wire ends should be smoothed to prevent injury. In use, the loop must be allowed to cool briefly after flaming before contacting cultures.

5.3. Pipettes → Needle‐Free Syringes, Glass Droppers, Spoons/Measuring Tools

In research laboratories, liquid handling, for example to create dilution series of microbes or reagents, is typically performed using micropipettes with single‐use disposable plastic tips or pre‐sterilised disposable plastic pipettes. However, multi‐use plastic—and, more sustainably, glass—graduated pipettes (typically ranging from 0.1 mL to 25 mL) are widely available from laboratory suppliers and online retailers. In addition, sustainable strategies for liquid transfer include the use of plastic syringes without needles, which are washable and durable, as well as glass droppers fitted with rubber bulbs (Figure 1f). For approximate measurements in non‐critical tasks, metal teaspoons or kitchen measuring jugs may also be sufficient. These alternatives are easy to clean, and some are sterilisable, reducing reliance on disposable plastic pipettes in educational settings where high precision is not required (Bayrau et al. 2025; Trusler et al. 2024).

5.4. Tubes → Small Glass Bottles With Lids or Cotton ‘Plugs’

Small glass containers with lids (e.g., spice jars or sample vials) can serve as effective substitutes for test tubes for both solid and liquid cultures. Their wide availability, reusability and compatibility with heat‐based sterilisation methods make them well suited to sustainable school laboratories (Freese et al. 2024). Where metal lids are unavailable or undesirable, a traditional laboratory alternative is the use of cotton ‘plugs’ (sometimes referred to as ‘dolls’), prepared from cotton wool wrapped in cotton gauze and fitted to the mouth of the container (Figure 1e). These plugs can be adapted to different diameters and, when properly fitted, help reduce contamination while allowing limited gas exchange.

5.5. Filter Paper/Wipes → Sterilizable Textiles and Reusable Filters

For routine cleaning and basic filtration, washable cotton cloths can replace paper towels and disposable wipes, provided they are thoroughly laundered and sterilised (e.g., using an autoclave or pressure cooker) between uses. In some cases, fine‐weave cotton fabric or reusable coffee filters can serve as substitutes for laboratory filter paper, particularly for non‐analytical tasks where precise filtration performance is not required.

5.6. Gloves → Reusable Household Gloves and Reinforced Hand Hygiene

In low‐risk activities involving non‐pathogenic microorganisms (e.g., well‐characterised probiotic bacteria), reusable rubber gloves that can be cleaned and disinfected between uses may be used in place of disposable nitrile or latex gloves, provided that appropriate risk assessment and supervision are in place (Figure 1f). Gloves should be washed with soap and water and then disinfected (e.g., with an alcohol‐based disinfectant) and inspected regularly for degradation or tears. Emphasising rigorous handwashing before and after handling materials further reinforces good hygiene practice and helps reduce reliance on single‐use plastics.

A summary comparison of conventional equipment/materials and their sustainable counterparts is provided as ‘Comparative overview: standard laboratory equipment/materials vs. environmentally sustainable alternatives for school Microbiology’ in Supporting Information document Table S1.

6. Accessible and Safe Sources of Microorganisms for BSL‐1 Teaching Contexts

Access to suitable microorganisms is pivotal for planning hands‐on microbiology activities in secondary education. Although the isolation and cultivation of microorganisms are central to microbiological learning, sourcing pure cultures from commercial suppliers can be cost‐prohibitive and may be subject to institutional or regulatory constraints in school settings. Accordingly, it is important to prioritise sources that are accessible, low‐cost, low‐risk and compatible with biosafety level 1 (BSL‐1) classroom work, while maintaining clear biosafety boundaries.

A practical alternative to purchasing strains is to use food‐associated and probiotic microorganisms as comparatively controlled, pedagogically rich inocula. These sources enable work with viable microbes commonly regarded as low risk in educational settings, while supporting learning about microbial diversity, fermentation and basic aseptic technique. Nevertheless, schools should conduct a simple risk assessment and comply with local requirements for handling, containment and disposal—even when organisms originate from consumer products—because cultures can vary and contamination can occur.

6.1. Dairy Ferments: Yoghurt (Lactic Acid Bacteria)

Natural yoghurt provides an accessible source of Lactobacillus delbrueckii subsp. bulgaricus and Streptococcus thermophilus —two well‐characterised lactic acid bacteria commonly used in dairy fermentation. When diluted in sterile water and inoculated onto simple media (e.g., milk agar), yoghurt cultures allow students to observe acidification, colony morphology and growth dynamics, offering a tangible connection between theory and practice (Pakpour and Hussain 2020; Kuzmenko et al. 2022).

6.2. Baker's Yeast: Saccharomyces cerevisiae

Commercial baking yeast is a readily available source of Saccharomyces cerevisiae , a widely used eukaryotic model organism. Its ease of cultivation in sugar‐based broths or glucose‐containing solid media makes it well suited to classroom experiments on fermentation, carbon dioxide production, respiration and eukaryotic cell structure (Marshall 2019; Chan et al. 2021). Bread or dough preparation can also be used as a low‐risk introductory demonstration, linking microbial metabolism to everyday food science.

6.3. Kombucha (SCOBY)

Kombucha contains a symbiotic culture of bacteria and yeast (SCOBY), often including acetic acid bacteria and yeasts. As a living consortium, it can support interdisciplinary activities that integrate microbial ecology and biochemistry (Applegate et al. 2019; Guzman‐Cole and García‐Ojeda 2022; Nyhan et al. 2022). Because microbial composition can vary between batches and sources, kombucha is best framed as a model for community‐based fermentation rather than a single, standardised culture.

6.4. Kefir (Milk/Water Kefir) and Ecological Value/Consortia

Kefir grains host interactive communities of bacteria (e.g., Lactobacillus, Streptococcus) and yeasts (e.g., Kluyveromyces), providing a strong platform for exploring symbiosis, niche partitioning and microbial interactions (Blasche et al. 2021). Citizen‐science approaches have used milk and water kefir fermentations in school and home contexts to support learning about microbial ecology and fermentation (Walsh et al. 2024). As with other fermentations, composition can be variable and should be treated as a feature of the model system rather than a limitation.

6.5. Fermented Vegetables (Sauerkraut/Kimchi)

Naturally fermented vegetables provide rich sources of lactic acid bacteria, including Lactobacillus plantarum and Leuconostoc spp. Liquid extracted from these foods can be plated directly, enabling exploration of fermentation ecology, succession and selective media use. Sauerkraut‐based activities have been used in interdisciplinary teaching laboratories to illustrate microbial diversity and biochemical processes (Hauptmann et al. 2025). Fermented Brassica foods also provide an entry point for discussing links between microbiology, health and environmental systems (Fijan et al. 2024).

6.6. Commercial Probiotics (Capsules/Sachets) as a Reproducible Source

A key aim in microbiology education is to reveal the ubiquity of microbes and our personal relationship to them, including the fact that we consume them every day, also to counter germophobia. Probiotic supplements (capsules, sachets or liquids) offer a well‐characterised, highly controlled, reproducible and safe source of beneficial microbes for educational microbiology. They also allow students to visualise the living microorganisms present in products they consume, bridging microbiology with everyday health and nutrition. Because availability, branding and formulations vary by country and may change over time, brand names should be treated as region‐specific examples and replaced with locally available equivalents; teachers should record product type, stated strains (and counts if provided), expiry date and storage/handling conditions to support reproducibility. Commonly studied organisms found in these products include:

• Saccharomyces boulardii (marketed locally as probiotic capsules or sachets in various regions, e.g., Ultra‐Levure, Floratil), a probiotic yeast with widely documented use in preventing antibiotic‐associated diarrhoea and modulating microbiota composition.

• Lactobacillus reuteri (available in drops or capsule formulations in many regions, e.g., BioGaia), noted for its ability to colonise the human gut and influence host physiology.

• Bifidobacterium and Lactobacillus blends (multi‐strain probiotic formulations, e.g., Bion3, VSL#3), commonly used in research and clinical contexts; these preparations illustrate the complex interactions within mixed probiotic formulations.

• Bacillus subtilis var. natto (natto starter cultures marketed in various regions, e.g., Startercultures ‘Nattō’ starter culture), a spore‐forming bacterium widely associated with food fermentation; it is robust and well suited for classroom discussions of sporulation, environmental persistence and fermented foods.

• Clostridium butyricum (present in certain probiotic formulations in some regions, e.g., Vitamatic ‘ Clostridium Butyricum ’), an anaerobe associated with butyrate production; it can support discussion of oxygen sensitivity and anaerobic metabolism, provided activities remain within the framework's defined biosafety boundaries and use controlled, low‐risk protocols.

When rehydrated in sterile water or, preferably, sterile isotonic saline (0.85%–0.9% NaCl) to minimise osmotic stress during rehydration, and then plated on general‐purpose media (e.g., nutrient agar) or selective media (e.g., MRS for lactic acid bacteria), these supplements enable students to culture and observe live, well‐defined strains in a classroom setting.

Fermentation is a globally shared microbiological practice, not a Western or laboratory‐only tradition. Teachers can leverage locally familiar fermented foods and starter cultures as culturally situated entry points into microbial growth, metabolism and community interactions (e.g., cereal fermentations such as ogi/akamu; maize beverages such as chicha; dairy, legume or vegetable ferments depending on the region). Such examples can support student engagement by connecting microbiology to everyday knowledge systems, while the framework's biosafety boundaries (containment and validated inactivation) remain non‐negotiable.

6.7. Higher‐Uncertainty Environmental Materials (Teacher‐Led, Sealed Observation Only)

In contrast to the sources above, environmental materials (e.g., soil suspensions or household moulds) contain microorganisms of unknown composition and should therefore be treated conservatively in school settings. Where such materials are used at all, they should be framed explicitly as teacher‐led demonstrations conducted under strict containment, with cultures kept sealed throughout incubation and observation, and with no subculturing or open handling after incubation. This distinction reinforces a core biosafety principle: unknown microbial communities must be treated as potentially containing pathogenic strains until proven otherwise.

Several familiar materials can still be used safely for observational purposes under these constraints:

• Overripe fruit skins (e.g., bananas or grapes): naturally colonised by wild yeasts and lactic acid bacteria; suitable for illustrating microbial presence and fermentation‐related concepts when handled with sealed vessels and prompt inactivation (Tournas and Katsoudas 2005).

• Bread mould (collected with caution): visible filamentous fungi can support teaching fungal morphology and sporulation; however, due to rapid sporulation and potential respiratory irritation, observation should be restricted to sealed containers and brief, supervised demonstrations, ideally in well‐ventilated conditions (CDC 2020).

• Diluted garden soil (handled with care): soil suspensions can reveal diverse microbial populations and support ecological discussion; however, because the composition is unknown, activities should be teacher‐led and contained, with sealed incubation and reliable end‐of‐session inactivation (Petersen and Chan 2020; Fatton et al. 2021).

In all cases, basic safety measures (sterile tools, clear labelling, sealed incubation, avoidance of opening incubated cultures and reliable end‐of‐session inactivation) should be treated as non‐negotiable classroom routines.

6.8. Pedagogical Note: Natural Variability of Sources and How to Use It Didactically

Many of the accessible sources suggested above—particularly fermented foods, kefir grains and kombucha SCOBYs—are inherently variable. Their microbial composition can differ markedly between starters, batches, households and production conditions (e.g., temperature, substrate composition, salt concentration, fermentation time and handling). Studies on kombucha starter cultures and reviews of kombucha fermentation consistently report substantial variation in SCOBY‐associated communities across samples and production contexts, underlining that ‘the source’ is rarely microbiologically identical from one preparation to another (Ben Saad et al. 2025). Similarly, kefir grains are well established as complex, mixed consortia whose community structure can vary across grains and settings, including geographically distinct origins and differing cultivation conditions (Alraddadi et al. 2023). Fermented vegetables also exhibit reproducible patterns of succession while remaining variable across household processes and recipes, making them suitable for comparing how conditions shape community dynamics and metabolic outputs (Tlais et al. 2022; Thierry et al. 2023).

Rather than treating this variability as a drawback, it can be used as a didactic strength to teach core concepts in microbial ecology, experimental design and scientific inference, for example because of the exceptional diversity of microbes and the metabolic redundancy inherent in this diversity, particular functionalities, such as those required for production of a standard kefir, may be provided by different combinations of species. Students can compare replicate cultures prepared from different sources (e.g., two yoghurt brands; homemade versus commercial sauerkraut; different kombucha starters) and record differences in colony morphology, growth rate, pH change, gas production or odour development. These observations provide an authentic basis for discussing selection pressures, succession and functional redundancy, and for explaining why complex biological systems may not yield identical outcomes even under broadly similar conditions. Making this explicit can help students appreciate that variability and diversity of responses to change is not a flaw but a characteristic of living systems (including human behaviour), and opens opportunities to discuss why biologically simplified systems, such as agricultural monocultures, may be more vulnerable to disease and environmental disturbance.

Methodologically, natural variability provides a practical context for teaching replication, controls and standardisation. Learners can define which variables should be held constant (e.g., incubation temperature, medium composition, inoculum volume and incubation time) and which can be intentionally varied, while documenting outcomes using simple shared protocols (e.g., incubation logs, photographic records and basic quantitative scoring of colony types). In parallel, schools may contrast variable mixed consortia with more standardised sources (e.g., labelled probiotic products), reinforcing how reproducibility requirements differ between ‘model’ systems and community‐based fermentations. Finally, citizen‐science approaches demonstrate that fermentation‐based microbiology projects can be implemented at scale while retaining educational value, using variability as an object of enquiry rather than a confound to be eliminated (e.g., Walsh et al. 2024).

7. Low‐Cost Culture Media From Accessible Ingredients: Design Logic, Standardisation and Exemplars

In contexts where access to commercial microbiological media is limited or cost‐prohibitive, the preparation of culture media from household and locally sourced ingredients can provide an effective, pedagogically sound and sustainable alternative (Estes et al. 2021). Beyond improving accessibility, this approach supports inquiry‐based learning by encouraging resourcefulness, ecological awareness and conceptual understanding of microbial metabolism. Incorporating natural indicators into media can enhance interpretation by allowing students to visualise metabolic changes (e.g., acidification) directly during growth.

7.1. Functional Components of Culture Media and Accessible (Household−/Retail‐Available, Food‐Grade) Substitutes

Preparing culture media in school settings offers three pedagogical advantages: (i) it enables nutrient composition to be aligned with the organisms and learning aims; (ii) it provides a cost‐effective route to practical microbiology without exclusive dependence on commercial media; (iii) it supports active learning by making microbial growth observable and discussable in terms of evidence, controls and variability (see media recipes); and (iv) it offers an accessible entry point to nutritional thinking (requirements, sourcing, and the link between resources and growth), reinforcing biological principles across organisms.

Low‐cost media built from widely available ingredients have also been validated in peer‐reviewed educational work, supporting their legitimacy as a classroom strategy rather than an improvised substitute (e.g., Scharfenberg and Marquardt 2015). School‐based media preparation can be framed as the assembly of a small number of functional components. Making these components explicit supports ‘fit‐for‐purpose’ formulation, transparent reporting and reproducibility across classrooms.

• Carbon/energy source: A readily metabolised carbon source can be provided by food‐grade sugars, selected to match the learning objective (general growth versus fermentation/differential readouts). Accessible options include table sugar (sucrose), glucose/dextrose powders (often sold for brewing or sports use) and lactose (commonly available as a brewing or supplement ingredient). When the goal is comparative inquiry, concentrations should be reported in g L⁻¹ and kept constant across conditions.

• Nitrogen source and growth factors: In conventional microbiology, peptones and tryptone supply peptides, amino acids and growth factors. In constrained school settings, the most consistent household−/retail‐available, food‐grade substitutes are hydrolysed protein preparations, particularly whey protein hydrolysate (unflavoured) because hydrolysis increases the fraction of short peptides available for microbial uptake. Plant‐derived alternatives include soy flour and other legume flours (e.g., chickpea), which can support growth but are more variable in composition and may require clarification (settling/filtration) for clearer plates. Where a yeast‐derived supplement is available, nutritional yeast/yeast extract–type products can provide B vitamins and additional growth factors; these should be reported by product type and brand because formulations differ substantially.

• Salts and buffering capacity: Most general media require only modest ionic strength. Non‐iodised table salt or sea salt can be used, but iodised salts and salts with anti‐caking agents can introduce variability.

• Vitamins and trace factors (optional): For enrichment or fastidious organisms, vitamins/trace factors may be supplied indirectly through yeast‐derived products (as above) or through small additions of food‐based extracts. Common trace factors include iron, magnesium and manganese. Because these inputs are chemically complex, they should be treated as optional enrichments and clearly documented.

• Gelling agent for solid media: For solid media, agar remains the preferred gelling agent. Importantly, agar can be sourced as food‐grade agar‐agar (E406) from local markets/cooperatives. Commercial ‘vegetable gelatin’ mixes marketed as agartina/agartine can also be used where locally available, but these products may contain agar‐agar blended with carriers (e.g., maltodextrin) and therefore vary in gelling strength; concentration optimisation may be required and should be reported by brand and dose. Gelatin is inexpensive but is limited by soft gels, melting near 37°C and potential liquefaction by gelatinase‐positive bacteria; it is therefore best reserved for low‐temperature demonstrations.

• Water quality is a frequent hidden source of variability. Where available, use distilled or reverse osmosis/deionised water (from laboratory suppliers or locally sold demineralised/deionised water for appliances or vehicle maintenance); otherwise, use tap water and record the source and any treatment (e.g., boiling/standing time) to support interpretation of variability.

These substitutions are intended to enable biosafe, educational, non‐diagnostic microbiology. Accordingly, the emphasis should be on transparent formulation, internal comparison (controls) and interpretive discussion rather than on replicating clinical or research performance specifications.

7.2. Media Design in Constrained School Settings: Functional Archetypes and Decision Criteria

Rather than reproducing the breadth of commercial catalogues, school‐based microbiology benefits from a small number of functional media archetypes that can be assembled from accessible ingredients and adapted locally. The key design decision is to define the instructional function of the medium—supporting broad growth, biassing community composition or generating a differential/indicator readout—and then to select the minimum components required to deliver that function with transparent constraints.

In practice, three archetypes cover most classroom investigations:

• General‐purpose (‘growth’) media: formulated to support robust colony formation and allow basic aseptic technique, enumeration and morphology comparisons. Here, the priority is consistency across batches, so ingredients should be specified precisely (including brands where variability matters), and formulations should be described as ‘LB‐like’ (a simple broth containing basic nutrients such as peptone/yeast extract and salt) when food‐grade substitutes are used.

• Selective or biassing media (ecology‐oriented): designed to shift competitive balance through pH, osmolarity or nutrient limitation, enabling discussion of environmental constraints and niche selection. These media are particularly aligned with Eco‐Microbiology because the ‘selection pressure’ is both mechanistic and conceptually teachable (e.g., acid biassing for fungi/yeasts; salt biassing for halotolerant microbes).

• Differential/indicator media (metabolism‐oriented): designed primarily to produce an interpretable phenotypic readout (e.g., acidification via carbohydrate metabolism), often by incorporating a built‐in pH indicator or by using an external indicator in parallel. In these cases, the medium is not judged by maximal growth yield but by clarity and reliability of the signal, and indicator choice should be justified by its pH transition range and classroom safety profile (see Section 7.3).

To ensure reproducibility and reduce ‘recipe drift’, each formulation should be reported using a minimum information template (water type, final concentrations, target pH and whether measured/estimated, sterilisation approach and intended use), with the complete set of recipes provided in Table S2 (‘Representative low‐cost media archetypes using accessible ingredients’). Extensions to additional organism groups can be guided by curated repositories (CCAP Media Recipes; ActinoBase Media Recipes; Microbe Online – Fungal Media), allowing teachers to scale complexity without inflating the core manuscript.

7.3. Natural Indicators Incorporated in Media (Red Cabbage/Anthocyanins)

Where rapid, visual readouts are pedagogically advantageous, pH‐responsive natural indicators can be incorporated into media or used in parallel as colorimetric reporters. Red cabbage ( Brassica oleracea var. capitata f. rubra) is rich in anthocyanins whose visible colour depends strongly on protonation state, providing a low‐toxicity and biodegradable alternative for classroom demonstrations and inquiry‐based investigations (Chigurupati et al. 2002; Abedi‐Firoozjah et al. 2022).

Beyond education, anthocyanin indicators underpin ‘intelligent’ food‐packaging concepts and freshness sensors, offering an authentic bridge between microbiology, chemistry and food technology. A classroom‐validated example is the red‐cabbage sauerkraut activity described by Linder et al. (2018), which uses anthocyanin colour shifts to track fermentation‐driven acidification while explicitly linking acid–base chemistry and microbial metabolism.

To support reliable classroom use, preparation should be standardised (plant mass‐to‐water ratio, extraction time/temperature, filtration) and the extract stored cold and protected from light; indicator performance is sensitive to pH and temperature and can drift across sessions (see ‘Red cabbage (anthocyanin) indicator incorporated in media: standardised protocol, storage, use and data recording.’ in Supporting Information document Box S2 for details). In classroom use, the indicator supports real‐time discussion of microbially driven acidification, particularly in lactic acid fermentation systems.

7.4. Other Plant Sources and Selection Rationale (pH Window)

To complement red cabbage anthocyanins, other edible plant extracts can be used as pH‐responsive colour reporters, enabling students to compare indicator performance and to relate pigment chemistry to observable signal behaviour. Suitable options include additional anthocyanin sources (e.g., hibiscus, berries, grape skin), betalain‐rich materials (e.g., beetroot), polyphenol mixtures (e.g., strong black tea) and curcumin (turmeric), selected primarily for their effective transition range and visual contrast in the chosen assay format.

Selection should be guided by four practical criteria: (i) expected direction and magnitude of pH change (choose an indicator whose transition window overlaps the anticipated shift); (ii) signal readability (contrast against agar colour/turbidity and under classroom lighting); (iii) stability and handling constraints (sensitivity to heat, light and storage time); and (iv) availability and standardisability (consistent sourcing and a preparation method that can be replicated across sessions). When students are invited to propose and justify an indicator choice (e.g., hibiscus vs. turmeric), the activity can strengthen experimental design ownership while reinforcing the link between chemical structure, pigment behaviour and interpretation of results.

In the Supporting Information, Table S3 summarises representative edible plant indicators, their approximate colour behaviour under acidic, near neutral and basic conditions, and their dominant pigment classes (Pandit et al. 2023; Perera et al. 2025). As with red cabbage, observed colours should be interpreted as qualitative to semi‐quantitative unless triangulated with an independent pH measurement; batch‐to‐batch variation is expected. When comparisons across groups are intended, users should prepare a simple in‐class reference (indicator solution alongside at least two known pH points) and document the extraction and observation conditions.

8. Biosafety, Disinfection, Sterilisation and Waste Management

This section summarises feasible, school‐appropriate procedures for handling microbial cultures safely before, during and after classroom activities. The approach is designed for low‐resource settings and emphasises good microbiological practice, clear operational routines and reliable inactivation of cultures prior to disposal. Although this manuscript prioritises low‐risk, food‐associated microorganisms commonly handled in BSL‐1 teaching contexts, a core principle is that any culture of unknown composition must be treated conservatively. An operational risk assessment and biosafety checklist, designed for BSL‐1 teaching contexts and intended as a printable pre‐lab gatekeeping tool, is provided in the Supporting Information (Table S4). However, checklists are essentially prompts; it is equally important to develop a culture mentality of laboratory safety based upon the precautionary principle and awareness of potential hazards.

8.1. Operational Definitions: Cleaning vs. Disinfection vs. Sterilisation

Clear distinctions between routine hygiene procedures are essential for safe microbiology teaching:

• Cleaning refers to the physical removal of dirt and organic material, typically using water and locally available cleaning agents (e.g., soap, detergents or other surfactant‐based materials). Cleaning reduces microbial load primarily by removal and improves the effectiveness of subsequent disinfection.

• Disinfection refers to the inactivation of many (but not necessarily all) microorganisms on surfaces or objects, commonly using chemical agents such as alcohol or sodium hypochlorite or equivalent alcohol‐based solutions and chlorine‐releasing compounds, where available.

• Sterilisation refers to the complete elimination or inactivation of all viable microorganisms, including bacterial spores. In school contexts, the most practical route to sterilisation is moist heat under pressure (e.g., a pressure cooker).

• Accidents happen; adopt the principles of containment and render harmless. If a spill of a liquid culture or a ‘plate’ of a culture falls on the bench/floor, clean up immediately, sterilise the material used to clean up and disinfect the surfaces that are affected. Wash hands scrupulously. Document the accident and the response, who was involved and the timeline of the response.

8.2. Core Principle: ‘Unknown = Potentially Pathogenic’

International guidance distinguishes risk group 1 (RG1) organisms—generally considered non‐pathogenic to healthy humans—from organisms of unknown risk. Many microorganisms used in school activities can be selected from low‐risk, food‐associated sources; however, a critical safety principle is that all unknown microorganisms should be treated as potentially pathogenic until proven otherwise. This distinction between known low‐risk sources and unknown or environmental isolates should be made explicit in educational settings (WHO 2004; CDC 2020).

In practice, this principle requires strict behavioural and procedural barriers, including teacher supervision, avoiding direct contact with incubated materials, keeping culture vessels sealed whenever possible and ensuring reliable inactivation of all cultures at the end of activities. Such routines support a biosafety culture that prevents accidental exposure and helps students understand scientific responsibility and ethical laboratory practice. Low‐risk microorganisms can be handled in basic educational laboratory settings without specialised containment equipment, provided that good microbiological practice is followed (e.g., Carlberg and Yeaman 2006; Emmert 2013; Byrd et al. 2019).

In this paper, exposure risk is understood primarily in terms of common contact/entry routes in classroom settings (hand‐to‐mouth, aerosols, splashes to eyes/skin), and is managed through behavioural rules, sealed handling and validated inactivation. The main hazards in a microbiology experiment are (i) ingestion—materials enter the mouth; (ii) inhalation—aerosols are breathed in; (iii) body surface (usually hands) contact with materials. Key issues are therefore avoidance of anything entering the mouth, of aerosols or direct contact with microbial samples—that is, only indirect contact with inoculation loops, etc. In practice this means an absolute ban on food/eating, drinks/drinking, putting fingers‐pencils‐pens‐anything else in mouths, and having skin disinfectants available on every bench. Aerosols are particularly problematic because they can disperse through shared airspace and exposure may occur without being immediately recognised. Any activity that disrupts liquid‐air interfaces, such as shaking or pouring microbial suspensions, can generate aerosols and must be carried out with particular care.

Core good practice in school microbiology includes:

• Washing hands before and after handling cultures.

• Disinfecting work surfaces before and after experiments.

• Avoiding mouth pipetting, ingestion or contact with open wounds.

• Clear labelling and secure containment of all culture vessels.

• Minimising aerosol generation and avoiding unnecessary opening of plates or jars.

• Ensuring that work involving unknown or environmental materials is teacher‐led and appropriately contained

Additional precautions for fungi and moulds: even in low‐risk contexts, environmental fungi (e.g., bread mould) may cause allergic reactions or respiratory irritation due to airborne spores (CDC 2020). Where such materials are used at all, activities should be restricted to sealed containers and brief, supervised demonstrations, ideally in a well‐ventilated area. Teachers may also consider face coverings to reduce spore inhalation risk, and cultures should be inactivated and discarded promptly after observation.

8.3. Chemical Disinfection: 70% Ethanol and Hypochlorite (Good Practice)

Chemical disinfection is an important supplementary control measure in school microbiology. It supports safe practice by reducing microbial contamination on benches, equipment and hands during activities and by enabling effective management of minor spills. However, disinfection does not replace sterilisation where complete inactivation of cultures or contaminated materials is required. Accordingly, chemical disinfectants should be used alongside moist heat sterilisation for end‐of‐session treatment of cultures and waste.

1. Ethanol (70%) is a rapid‐acting disinfectant that disrupts membranes and denatures proteins (McDonnell and Russell 1999). It is suitable for routine disinfection of benches and tools (e.g., scissors, forceps and handles) and can be used during class to maintain clean working conditions due to its quick evaporation and low residue. Handwashing with soap and water remains the preferred approach for hygiene, but alcohol‐based products can be used when sinks are not immediately accessible.

2. Sodium hypochlorite (household bleach, diluted) is a broad‐spectrum disinfectant that oxidises essential cellular components and can inactivate a wide range of bacteria, fungi and viruses (e.g., Köhler et al. 2018). For classroom use, standard household bleach (often ~5% sodium hypochlorite) may be diluted to a working solution (e.g., ~0.5% available chlorine), suitable for decontaminating used culture vessels and managing spills. Solutions should be clearly labelled, used in well‐ventilated areas and prepared regularly according to product guidance. Because bleach can irritate skin and eyes, protective measures (e.g., gloves, eye protection and ventilation) are required.

Where cultures and contaminated materials must be rendered non‐viable prior to disposal, moist heat sterilisation (Section 8.4) should be used whenever feasible. Chemical disinfectants can complement this workflow by supporting routine surface hygiene and safe spill response.

8.4. Moist Heat Sterilisation (Pressure Cooker)

Moist heat under pressure is the most reliable and sustainable method for eliminating viable microorganisms, including bacterial spores. A domestic pressure cooker can approximate autoclave conditions by reaching temperatures of approximately 121°C under pressure. In school contexts, a practical cycle is 15 min once full pressure is reached, which can be used to inactivate cultures in media, sterilise reusable glassware and treat contaminated materials prior to disposal (CDC 2020).

This approach is advantageous because it is effective, leaves no chemical residues and reinforces key microbiological concepts (protein susceptibility to denaturation, microbial resistance, sterilisation parameters and safe waste handling). After a cycle, materials should be allowed to cool fully before handling and containers should remain sealed until they can be safely cleaned or disposed of.

8.5. Waste and Inactivation: Safe Workflow From ‘End of Experiment’ to Disposal

A clear end‐of‐session workflow is essential to ensure that no viable organisms are released into the environment and that classroom routines remain consistent and safe (Emmert 2013; Byrd et al. 2019; CDC 2020). A practical school workflow includes:

1. Seal and label: At the end of the activity, close and seal culture vessels (e.g., plates, jars, tubes) and label them clearly (date, contents/source, class/group).

2. Inactivate cultures: Preferably inactivate all cultures by moist heat sterilisation (pressure cooker). Where immediate sterilisation is not possible, keep cultures sealed and securely contained; use chemical disinfectants for surface decontamination and spill management and sterilise the cultures as soon as practicable.

3. Cool and contain: Allow sterilised items to cool completely before handling. Avoid opening culture vessels unless necessary for cleaning or disposal.

4. Separate waste streams: Keep reusable glassware separate from single‐use items. Treat liquid cultures and solid materials as contaminated until inactivated.

5. Dispose responsibly: Once inactivated, dispose of waste according to local requirements and school policies. Reusable items should be cleaned and prepared for the next use.

Figure 2 summarises a school‐appropriate decision pathway from the end of an experiment to disposal. It highlights key control points—sealing and labelling, inactivation (preferably via pressure cooker) and segregation of reusable versus disposable items—providing a simple visual guide for implementing consistent biosafety routines in classroom microbiology.

FIGURE 2.

Inactivation and Waste Management of Microbial Materials in Secondary Education Laboratories (BSL‐1 context).

9. School Implementation Guide: Planning, Practice and Assessment

This implementation guide translates the approaches described in this article into feasible classroom practice. It is designed to be adaptable to different school contexts, including settings with limited equipment, constrained timetables and variable access to consumables. The emphasis is on safe, low‐cost microbiology that remains pedagogically rigorous through clear routines, appropriate controls and structured documentation. Before any activity is run, instructors should apply the operational checklist (Supporting Information, Table S4) to classify sources/materials and to define permitted manipulations and end‐of‐session inactivation.

9.1. Planning and Preparation: Aligning Objectives, Resources and Biosafety

Effective implementation begins by aligning curricular objectives with resource constraints and biosafety boundaries. Teachers should define a small set of intended learning outcomes (e.g., students can perform a basic aseptic transfer following a checklist without spills; calculate and justify a simple dilution series; record a complete logbook entry including controls and deviations; interpret results using negative controls and stated limits of inference; and select/justify a frugal substitution using the decision criteria in Section 3.3.), then select an activity that fits the timetable (including incubation time and a defined end‐of‐session inactivation route).

Before implementation, educators should:

1. Define learning goals (e.g., microbial growth as evidence; fermentation‐driven acidification; the role of temperature and oxygen; what can and cannot be inferred from colony appearance and colour change).

2. Select an appropriate microbial source. Prefer controlled, food‐associated sources suitable for BSL‐1 teaching (e.g., plain natural yoghurt cultures, probiotic preparations) and avoid open handling of unknown/environmental materials beyond teacher‐led, sealed observation.

3. Secure an inactivation and disposal pathway before cultures are generated (preferably moist heat under pressure; see Section 8).

4. Plan containment and transport: label all vessels before inoculation; use sealed incubation; use secondary containment (tray/box) for transport and storage.

5. Pre‐validate incubation conditions (temperature stability and duration) and document them as part of the activity record.

6. Trial each experiment in advance under local conditions before implementing it with students. This increases the likelihood of success and allows the teacher to anticipate typical outcomes (including variability), identify likely points of failure (e.g., contamination, temperature instability, weak indicator response) and refine timings, controls and documentation requirements. If outcomes are inconsistent, one factor at a time should be revised and documented so that deviations become part of the learning about the interactive process of scientific sleuthing, evidence and uncertainty.

9.2. Classroom Routines That Increase Rigour and Reduce Contamination

To strengthen reproducibility and maintain biosafety, the following routines should be treated as minimum classroom standards:

• Bench disinfection and hand hygiene before and after work; clear ‘no eating/drinking’ and ‘no face touching’ rules.

• Sealed incubation and sealed observation: cultures remain closed during incubation and observation; students record results without opening plates/jars after incubation.

• Clear labelling (source, dilution, group, date, incubation condition) and basic documentation (time, temperature, container type and any deviations).

• Use of controls for interpretability (Section 9.4). Unexpected growth in controls should be treated as evidence about technique/contamination rather than as ‘failure’.

• Accident prevention and responses (spill plan, teacher‐led escalation and post‐incident cleaning/inactivation steps).

• Laboratory logbooks (recommended minimum standard): date/session; aim; microbial source/product details; method summary + critical parameters (time, temperature, volumes/dilutions); controls; observations/results (incl. photos); deviations/incidents; interpretation + limits of inference; and confirmation of end‐of‐session inactivation/waste route. Teachers may assess logbooks using a simple rubric (completeness, accuracy and reflection on deviations).

9.3. Model Activity (Overview)

A worked exemplar illustrating implementation of the framework in a secondary classroom is provided in the Supporting Information (Protocol S1). The exemplar uses yoghurt‐associated lactic acid bacteria cultured on milk‐based media under reduced oxygen exchange to demonstrate substrate–environment interactions, sealed incubation/observation and explicit end‐of‐session inactivation following the workflow in Section 8 and Figure 2. Detailed materials, media preparation (including milk proportions), stepwise procedures and minimum documentation standards are provided in the Supporting Information to support local adaptation without overloading the main text.

Pedagogically, this exemplar is intended to illustrate how frugal microbiology approaches can support both exploratory observation and question‐driven experimentation. Initial observations of microbial growth patterns may prompt investigable questions that guide experimental design; for example, students might examine how pH acts as a selection pressure by plating soil dilutions on media adjusted to contrasting pH values. Although the worked exemplar provided here focuses on one model activity, teachers may draw on curated (and evolving) classroom protocols hosted on the EduBiota platform (https://www.edubiota.com), as well as established practical guides and repositories that document school‐appropriate microbiology techniques (Grainger and Hurst 2016; Burdass et al. 2016; Lammert 2006; American Society for Microbiology, n.d.).

9.4. Controls and Comparisons (Minimum Set; Strengthens Inference)

Include at least two controls to improve interpretability and to model good scientific practice:

• Uninoculated medium control (incubated alongside samples): checks contamination introduced during media preparation/pouring.

• Air‐exposed control (briefly opened, then sealed and incubated): illustrates airborne contamination, reinforces aseptic technique and reveals what result non‐target organisms can create in the context of the imposed experimental conditions.

• Optional comparative design (recommended): oxygen condition comparison. Incubate one set in a sealed secondary container (reduced oxygen exchange) and another with standard incubation (no secondary sealing). This tests the role of oxygen availability and typically increases engagement and interpretive depth.

9.5. Optional Extensions (Inquiry‐Friendly, Low Additional Cost)

Extensions allow differentiation by age level and time available:

• Field extension (observation‐first): add a short excursion to document microbial habitats and collect contextual data (e.g., temperature or pH gradients; simple transects through compost heaps using basic thermometers; optional pH screening of soils or waters using strips or low‐cost metres), and, where available, portable/home‐made microscopy for direct observation (McGenity et al. 2020; Archer et al. 2025; IMiLI_Mic). Any environmental samples brought back for follow‐up work should be treated as unknown and remain within the framework's sealed, teacher‐led boundary with validated end‐of‐session inactivation and waste workflow defined in Figure 2.

• pH as selection pressure (ecology‐oriented): adjust the medium pH to contrasting values (e.g., acidic vs. near‐neutral) and compare colony diversity (morphotypes) and/or indicator readouts. Use this to discuss why diversity often peaks near neutrality and connect to real‐world issues such as soil acidification (e.g., acid rain) and ocean acidification.

• Osmotic stress (salt) effects: compare standard medium versus medium with added salt (within safe, food‐grade ranges) to illustrate selection for halotolerant microbes and to link to food preservation, fermentation and environmental constraints.

• Inoculum/dilution effects (evidence quality): compare two dilution levels of the same source to make ‘countability’, crowding and morphology visibility explicit; use this to discuss what counts as interpretable evidence and why dilution‐to‐extinction supports observation.

• Temperature effects: compare room temperature versus warm incubation (ideally 37°C–42°C; otherwise use a near‐body‐temperature proxy); discuss relevance to enzyme activity and growth kinetics.

• Substrate effects: compare milk‐based medium versus a general‐purpose nutrient rich ‘LB‐like’ school medium; discuss ecological fit and selective bias.

• Indicator‐based interpretation: compare media with versus without built‐in pH indicator; discuss qualitative versus semi‐quantitative inference and the need for simple reference anchors.

• Source comparison: compare two yoghurt sources (where available), or yoghurt versus a labelled probiotic; link outcomes to variability, standardisation and reproducibility.

9.6. Assessment: Prioritising Scientific Practice Over ‘Getting the Expected Result’

Assessment is most robust when it targets scientific practice and reasoning rather than ‘correct’ (textbook/expected/desired) outcomes:

• Lab notebook requirements: labelled photographs, incubation log (time/temperature/condition) and a short results table (growth/no growth; colony descriptors; indicator outcome).

• Interpretation prompts: What do controls tell you? Which variables were controlled? What is evidence versus interpretation? What limits your inference?

• Simple rubrics: aseptic handling behaviour, documentation quality, appropriate interpretation of controls and ability to articulate uncertainty and variability.

• Conventional assessment formats can also be used where aligned with local curriculum: (i) a short lab report (aim, method, results, discussion, limitations/uncertainty and biosafety/waste route reflection); and/or (ii) a poster or short oral presentation summarising the question, controls, observations, interpretation and the frugal substitution rationale (Section 3.3). These formats assess scientific communication and reasoning rather than ‘getting the expected result’.

10. Limitations and Considerations for Scale and Transfer

While these strategies broaden access to practical microbiology in resource‐limited schools, important limitations emerge when scaling across contexts. Even when the substitution criteria in Section 3.3 are met, the points below stress‐test the framework by clarifying expected sources of variability and the limits of inference, and the distinction between within‐class reproducibility (a primary goal) and cross‐school comparability (not a primary aim of the framework).

10.1. Precision, Reproducibility and Variability of Improvised Materials

Approaches based on non‐standardised materials (e.g., domestic containers, improvised incubators, handmade inoculation tools) are intentionally designed for accessibility and sustainability, but they may reduce experimental precision and reproducibility relative to conventional laboratory practice (Baker 2016; National Academies of Sciences, Engineering, and Medicine 2019). Temperature instability in improvised incubators, variation in container geometry and inconsistencies in handmade loops or transfer volumes can introduce variability in microbial growth rates, colony morphology and transfer efficiency. In exploratory and introductory activities this variability is often acceptable—and can itself be used didactically—yet it limits strictly quantitative comparisons when such comparisons are attempted (e.g., across groups or schools, as is desired in citizen science projects) unless protocols include clear standardisation steps (e.g., documenting incubation temperature, timing, inoculum dilution and container type).

10.2. Variability in Probiotics and Fermented Products and Implications for Comparability

Although classroom activities can prioritise low‐risk, food‐associated microorganisms typically consistent with risk group 1 (RG‐1) contexts, microbial composition and viability may vary across batches and products. Manufacturing processes, formulation updates, storage conditions and time‐to‐expiry can influence the presence and dominance of specific organisms in both fermented foods and commercial probiotics. This variability does not necessarily compromise safety, but it can affect experimental consistency. Because the framework prioritises safe participation and within‐class replication over inter‐institutional standardisation, such variation is usually acceptable in inquiry‐focused teaching. Where tighter standardisation is desired (e.g., within‐class comparisons across groups or across successive cohorts), educators may mitigate variability by selecting more standardised sources (e.g., labelled probiotic products), recording product details (brand, batch/lot where available, expiry date) and using consistent preparation steps (dilution, inoculation method, incubation conditions), alongside explicit controls. In this sense, variability is treated not primarily as a threat to cross‐site comparability, but as a parameter to be documented and discussed as part of scientific reasoning.

10.3. Dependence on Supervision and Biosafety Literacy

The effectiveness of biosafety routines depends on correct execution, consistent supervision and a shared understanding of good microbiological practice in the classroom. Domestic tools (e.g., pressure cookers, alcohol/spirit lamps) can be appropriate substitutes, but only when used with clear protocols, appropriate safety precautions, explicit classroom instruction and reliable end‐of‐session inactivation workflows. Inadequate supervision, poor labelling/containment or incomplete inactivation may increase exposure risk and undermine biosafety culture. Teacher training and rehearsal are therefore critical, including piloting activities in advance, clarifying non‐negotiable classroom rules (sealed cultures; no opening after incubation; controlled disposal) and ensuring that disinfection supports—rather than replaces—sterilisation when complete inactivation is required. Where feasible, schools may also benefit from a rapid‐access mentoring/consultancy contact (e.g., a nearby university, research centre, public health laboratory or professional society contact person) to support timely troubleshooting and situated biosafety decision‐making during implementation.

In most national curricula, the framework is best positioned within upper secondary biology or integrated science courses (typically ages 15–18), where students are introduced to microbiology, cell biology, metabolism or health and environmental science. Selected elements (e.g., fermentation, microbial growth observations or hygiene‐focused investigations) can also be adapted for lower secondary levels (ages 12–14 and below), under closer supervision and with simplified procedures. Implementation requires teacher supervision during all practical sessions involving heat sources or sterilisation and is most appropriate within structured laboratory blocks rather than unsupervised project work.

10.4. Minimum Infrastructure and Local Legal Constraints

Successful transfer also assumes the availability of minimum infrastructure, such as access to water for handwashing and cleaning, basic workspace that can be disinfected, and (in many cases) reliable electricity for incubation or safe lighting. In some school environments these prerequisites may remain limiting. In addition, regulatory and institutional constraints differ across countries and school systems: local rules may restrict the handling, storage, transport or disposal of microbial cultures—even those regarded as low risk—or may prescribe specific waste‐management pathways. Schools should therefore align all activities with local requirements and institutional policies, including those governing chemical disinfectants and waste disposal, and should ensure that implementation remains feasible within timetable constraints and local curricular requirements (plural) and learning outcomes.

Where feasible, a rapid‐access mentoring/consultancy contact (e.g., a nearby university, research centre, public health laboratory or professional society) can support timely troubleshooting and situated biosafety decisions; over time, such interactions can also be consolidated into shared troubleshooting notes that improve transferability and support teacher agency.

Despite these limitations, the approaches presented remain valuable for supporting meaningful, hands‐on microbiology in diverse settings. They are best treated as complementary strategies whose effectiveness is maximised when aligned with local biosafety guidance, teacher agency, and a clear understanding of where qualitative, inquiry‐focused learning is the primary goal versus where tighter standardisation is required.

11. Conclusion

The integration of microbiological investigations into secondary education is still frequently constrained by technical limitations, uneven infrastructure, biosafety requirements, time constraints and pedagogical barriers. This manuscript shows that these constraints can be mitigated through intentional instructional design and a pragmatic Eco‐Microbiology approach that safeguards scientific rigour while reducing costs and minimising environmental impact.

By prioritising reuse, local adaptation and low‐impact substitutions for conventional equipment and consumables, schools can implement meaningful, hands‐on and engaging practical activities while maintaining clear biosafety boundaries. To this end, the article presents an integrated, auditable framework for low‐cost microbiology using locally available materials and reusable solutions, supported by decision criteria and end‐of‐session inactivation and waste workflows suitable for school settings.

Beyond feasibility, the approach strengthens educational equity by enabling relevant practical experiences across diverse school contexts, including resource‐limited settings. By connecting microbiology familiar materials (e.g., fermented foods, probiotics, natural indicators), it can increase student relevance and support inquiry‐based learning. It also promotes interdisciplinarity by linking biology with chemistry (pH, indicators, metabolism), physics (temperature control, microscopy) and environmental science (materials, reuse, waste) and provides a platform for developing scientific citizenship through explicit attention to responsible practice, biosafety and environmental stewardship (Timmis et al. 2024).

Eco‐Microbiology also offers an age‐appropriate setting to foster critical thinking and systems thinking through scientific practice: distinguishing observation from interpretation, using controls to strengthen inference, articulating uncertainty and measurement limits and revising decisions when results are ambiguous (Timmis et al. 2025). Where feasible, rapid‐access mentoring arrangements with nearby universities, research centres or professional societies can strengthen teacher agency by supporting troubleshooting and situated biosafety decision‐making.

Finally, by making resource‐aware decision‐making and process discipline explicit, the framework helps connect foundational microbiology to emerging bioeconomy sectors, where biomanufacturing is increasingly framed within circular‐economy transitions (Takors 2025). Wider adoption will depend on teacher agency and institutional support; we recommend piloting activities in advance and maintaining simple documentation, appropriate controls and non‐negotiable close‐out routines to sustain safe, rigorous practice in school microbiology.

Author Contributions

Lara Amorim: conceptualization, investigation, writing – original draft, writing – review and editing, methodology, visualization. Kenneth Timmis: validation, writing – review and editing.

Funding

This work was supported by Fundação para a Ciência e a Tecnologia (FCT), 2024.04293.BDANA.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Box S1 Minimum QC for handheld pH measurement (school labs).

Box S2 Red cabbage (anthocyanin) indicator incorporated in media: standardised protocol, storage, use and data recording.

Protocol S1 Model activity: culturing yoghurt lactic acid bacteria on milk‐based, low‐cost media under reduced oxygen exchange.

Table S1: Comparative overview: standard laboratory equipment/materials versus environmentally sustainable alternatives for school Microbiology.

Table S2: Representative low‐cost media archetypes using accessible ingredients.

Table S3: Edible plant‐based pH indicators for classroom use: approximate colour transitions and dominant pigment class.

Table S4: Operational risk assessment and biosafety checklist for school microbiology (BSL‐1 teaching contexts).

Table S5: Transferable ‘life lessons’ communicated through practical school microbiology (BSL‐1 teaching contexts).

Data Availability Statement

Data sharing not applicable to this article as no datasets were generated or analysed during the current study.