OpenCell: Covid-19 Testing Laboratories in Shipping Containers

This living document is both a report of our ongoing work and a how-to guide for adapting and replicating the project. We will be updating each section as we progress, gather more data and characterise new techniques and methods.

If you want to contribute follow this link.
#ALL CONTRIBUTORS: Change ‘Editing’ to ‘Suggesting’ in the top right before contributing changes. Or, add contributions as comments.#

Converting a shipping container to BSL-2 safety level

Below information will provide you with all the necessary information to produce a portable BSL2 container lab including H&S and standard procedures.

Definition of BSL-2 category and useful guidance/resources

Definition BSL-2 -Biosafety level 2: A level of biosafety considered appropriate for agents that can cause human disease, but whose potential for transmission is limited.

World Health Organisation: Laboratory biosafety manual
Management and operation of microbiological containment laboratories

Container Type and Size

Type: Cargo container
Size: 30ft length, 8ft width

Type: Portacabin
Size: 32 ft length, 10 ft width

Requirements and modifications

The most affordable and time-sensitive way to purchase a converted shipping container is to reach out to a local producer with below requirements and attached technical plans.

  • 40 ft shipping container (used or new)
  • Insulation: Closed Cell Foam Insulation
  • Wall and ceiling cladding: PVCu Wall Cladding
  • Flooring: Commercial or Industrial PVC/Vinyl Flooring
  • Doors and windows:  2x UPVC Patio Doors (2200W x 2200H)
  • Partition wall with door


  • 1 External IEC60309 32A 240V Connection, 
  • 1. Internal RCD Consumer Unit, 
  • 4 1.5m Double LED Diffused Ceiling Lights c/w Switch, 
  • 5 Double 13a Sockets 
  • 1  Fused spur for over sink water heater
  • 1 Emergency light


  • Water supply and waste
  • 1 Stainless Steel Sink
  • 1 Undersink Water Heater (optional)
  • 1 Macerator / Grey Water Pump (143FV) (depending on local drainage)

Air conditioning (optional): 

  • 1 Wall Mounted Air Conditioning Inverter Unit 12000BTU

Interior Fit-Out

Link to interior and fit-out inventory.
Below are suggestions. In case you are using different products please read section "Adequate materials for the fit-out of a BSL lab".

Adequate materials for the fit-out of a BSL lab

Materials requirements for interior and fit-out

  • Smooth Surface
  • Washable
  • Chemical Resistant 
  • Non-Porous

List of suitable materials:

  • Stainless Steel
  • Polypropylene
  • Epoxy Resin
  • High-Pressure Laminate (e.g. Trespa)
  • Laminate Covered MDF (for shelving, cupboards etc. but not benchtops)

Materials NOT to use:

Wood, MDF, materials that a porous or can chip

Single Laboratory Overview

Below showcase the layout and workflow to achieve 2400 tests in 24h.

Lab Layout for 2,400 tests in 24h

Workflow to achieve 2,400 tests in 24h

Below spreadsheet showcases the workflow to achieve 2,400 tests in 24h. Stations will have to run in parallel to optimize the outcome. Headline showcases 24h in minutes.

Logging and Tracking Software

A digital sample management system has been built to track samples through the testing process, analyse and log test data, and securely communicate test results to the patient and/or healthcare provider.
Samples are tracked with unique IDs linked to a printable QR code (Fig. 6). During sample handling (Station A), sample QR codes are scanned and assigned to a test run with a unique run ID. At Station C, qPCR data is uploaded into the sample management system, and test results (i.e. positive, negative or inhibitory) are automatically generated. Results are communicated using the secure FHIR (Fast Healthcare Interoperability Resources) protocol, enabling data integration with NHS or international healthcare system architectures. The system is built using Ruby-on-Rails 5.2 on Ruby 2.4, with a PostgreSQL server to persist data, and operates on a cloud platform. Technical details and resources are found on our GitHub (ref: CoVid19/opencell-testing).

Scenarios and usecases

The modular and flexible nature of the CONTAIN system would allow it to be used in a variety of conditions. In this section we discuss three possible scenarios highlighting CONTAIN flexibility.

University Student and Staff Testing

Universities seeking to resume operation while ensuring the safety of their staff and student population will require regular testing capability. For a large, multi-campus university comprising for example 10,000 staff and 30,000 students, a full-time operation of 10,000 tests per week allows every individual to be tested monthly or more frequently if we consider the majority working from home (Fig. 7). Combined with selective quarantining, this capacity would allow a university to safely resume relatively normal teaching and research practices. Commitment to supporting the health and work of the student body also gives greater peace of mind to the significant overseas student populations who currently face additional challenges in travel, healthcare and accommodation. Two dedicated CONTAIN units can meet a 10,000 test per week capacity, with their mobility enabling them to operate at multiple campuses and accommodation sites throughout the week.

Augmenting existing medical infrastructure

Augmentation of existing healthcare infrastructure to meet the surge in demand for SARS-CoV-2 testing requires rapid deployment of suitable (BSL2+) laboratory space. CONTAIN units can be quickly transported to hospital sites and require only a car park area and utilities connection (electricity, water) to be operation-ready. The addition of a dedicated, separate facility enables a hospital to increase the scale of its testing without impacting other essential in-house laboratory work and maintains the safety of the main hospital building (Fig. 8). The flexibility of OpenCell’s CONTAIN system allows hospitals to integrate the unit with their own testing process, existing reagents, waste management system, and/or workforce. CONTAIN units can be transported overseas using standard shipping infrastructure, and additional units may be deployed or moved rapidly between hospitals to react to new outbreaks or surges.

Large Scale implementation: 39 container facility for mass-testing

Large cities make a good logistical case for large centralised testing facilities. In this scenario, we take advantage of the stackability of shipping containers and the modular nature of CONTAIN to envisage a multi-container testing centre [Fig. 9]. The facility, constructed of 39 containers has the potential to run up to 72,000 tests per day at full capacity, and provide office, storage and utility space for the running of the testing centre. This facility could be built rapidly and economically to serve a spike in COVID-19 cases. Once the spike of cases has passed, the facility could then be resourcefully separated back into individual containers and distributed around the country to act as local testing facilities and follow local outbreaks.

Standard Health & Safet Operating Procedures

Below will cover the necessary health and safety procedures you will have to set in place.

Health and safety documents

Health and safety procedures for personal protection

  • Labcoat must be worn at all times for work in the laboratory.
  • Appropriate gloves must be worn for all procedures that may involve direct or accidental contact with blood, body fluids and other potentially infectious materials. After use, gloves should be removed aseptically and hands must then be washed.
  • Personnel must wash their hands after handling infectious materials before they leave the laboratory working areas.
  • Safety glasses, face shields (visors) or other protective devices must be worn when it is necessary to protect the eyes and face from splashes, impacting objects and sources of artificial ultraviolet radiation.
  • It is prohibited to wear protective laboratory clothing outside the laboratory, e.g. in canteens, coffee rooms, offices, libraries, staff rooms and toilets.
  • Open-toed footwear must not be worn in laboratories.
  • Eating, drinking, smoking, applying cosmetics and handling contact lenses is prohibited in the laboratory working areas.
  • Storing human foods or drinks anywhere in the laboratory working areas is prohibited.
  • Protective laboratory clothing that has been used in the laboratory must not be stored in the same lockers or cupboards as street clothing.
  • A maximum of two people are allowed in the laboratory at any time.

Handling and disposal procedures for contaminated materials and wastes

It is the responsibility of all personnel to ensure the safe and correct disposal of all wastes produced in the course of their work. It is essential that the appropriate disposal procedures given below are strictly adhered to.

Waste disposal streams
The types of waste generated within the laboratory can be categorised as:  

Sharps waste consists of all contaminated and non-contaminated items capable of piercing the skin and includes lancets, needles, scalpels, sutures and contaminated disposable glassware. Be careful when handling sharps and resultant waste and dispose of in designated sharps bins. Never use the sharps bin for the disposal of any other type of waste. Waste needs to be autoclaved before picked up by medical/biological waste collectors.

Container to use: Sharps Bin
Wasteflow: Sharps bin -> Autoclave -> Pick-Up biological waste service -> Landfill

Clinical waste includes all other contaminated material (e.g. swabs, examination
gloves, specula,, etc.) and yellow clinical waste bags are available for the
disposal of this type of waste. Do not dispose of general waste in these bags as it leads
to an unnecessary increase in waste disposal costs. Waste needs to be autoclaved before picked up by medical/biological waste collectors.

Container to use: Biohazard cardboard container
Colour code for bin and bag: Yellow
Wasteflow: Sharps bin -> Autoclave -> Pick-Up biological waste service -> Landfill

General waste
Examples of general waste are packaging from consumables and paper towels and general waste bins are available for this type of waste. Under no circumstances should any sharps or clinical waste be disposed of in the general waste bin.

Container to use: Standard bin
Colour code for bin and bag: black
Wasteflow: Pick-Up standard waste service -> Landfill

Liquid waste
All liquid waste must be stored in leakproof containers with a screw- top or other secure lid and labelled appropriately with content. Snap caps, mis-sized caps, parafilm and other loose-fitting lids are not acceptable.

Sending and receiving infectious material procedure

Coming soon.

Testing and maintenance procedure

Coming soon.

Emergency procedures

Coming soon.

Lab cleaning procedures

Coming soon.

Station A: Logging and Plateing

In Station A, samples are unpacked and logged with a barcode scanner. Then 240 μL of each lysed sample is pipetted into a 2 mL 96 well deep well plate. Once a plate has been filled with 92 samples, the sample plate is passed to Station B.


Below equipment is for a single container lab  Station A capable of running 2400 tests in 24h.
Link to equipment inventory list.

  • 1 BSL2 Cabinet
  • 1 Benchtop Autoclave  (if used as a standalone lab)
  • 1 PC with internet connection for logging
  • 1 Ice Bucket
  • 1 Pipettes (which ones)
  • 1 Goggles
  • 1 Labcoat


Station B: RNA Extraction

Using Bomb Bio magnetic beads (link to page and protocol, plus original RNA extraction protocol we modified and validated first)

If viral samples have been inactivated with a lysis buffer in station A this work can be safely undertaken at containment level 2.  At 21 ± 2 °C (70 ± 4 °F) and 60 ± 10 % humidity.


Below equipment is for a single container lab  Station A capable of running 2400 tests in 24h.
Link to equipment inventory list.

  • 4 OpenTrons OT2
  • 4 OT2 Magdeck (OpenTrons)
  • 4 P300 8-channel Pipettes
  • 1 Undercounter Fridge (manual defrost!)
  • 1 Weighing scale
  • 1 Weighing boat set
  • 1 PC (same like for station A)
  • 1 Ice Bucket
  • 1 Pipette (handheld) 1ml multichannel (ethanol and IPA),
  • 1 Pipette (handheld) 20ul single channel (ONLY used at Station B)
  • 1 Goggles (same like for station A)
  • 1 Labcoat (same like for station A)


Prior to Station B

  • Put on PPE - lab coat, and  gloves.
  • Samples from station A should be in 96 deep well plate. Each well should hold   240ul of  inactivated sample in a guanidine isothiocyanate based lysis buffer.


  1. Check all necessary PPE available
  2. Check Inventory for permanent items
  3. Fetch reagents (beads and nuclease-free water in fridge, IPA and EtOH in rack opposite station)


  1. Soapy water - clean every surface, pipettes, self, gloves, lab coats
  2. Clean workspace (10% bleach)
  3. Clean desk with RNAseZap

OpenTrons Set-Up

Reagent setup [2x 96 deep well plates]

  1. Add 1 ml EtOH (1 ml per well) into all wells in a 96 deep well plate (slot 6), using manual 8-channel 1ml pipette. [Filter tips in slot 3 are used exclusively for EtOH and reused. Automation code maps tip → EtOH well → sample, such that no cross-sample contamination occurs during washing cycles.]
  2. Add reagents to deep well plate (slot 8), using manual 8-channel 1 ml pipette (IPA) and manual single channel 20 ul pipette (otherwise):
  • IPA in columns 8-12 inclusive (2 ml per well)
  • Silica-coated magnetic beads in column 7 (0.5 ml per well)
  • Nuclease-free water in column 5 (0.5 ml per well)

Set up OT2

1. Connect via laptop (wifi or ethernet)
If first run, calibrate deck

2. Load automation protocol code

3. Follow calibration instructions for each piece of equipment (see diagram below).

[2x 96 deep well plates with reagents (slot 8) and EtOH (slot 6)]
[6x 96 filter tip racks 200ul (slots 2-5 inclusive, 7, 11)] #NB MAY NEED EXTRA RACK IN 10
[1x 96 well plate (slot 1)]
[MagDeck (slot 9)]
[Samples in 96 deep well (slot 9)]

Calibration can also be done using a “dummy” set of tips and plates which are reused for each calibration. This lowers the risk of contaminating fresh plastics.

4. When calibrated, virtual deck should appear as above.

5. Remove plasticware and clean OT2 deck and pippet with RNAseZap. (if using a dummy set for the calibration step replace the OT2 with the intended plasticware after cleaning)

6. Run protocol.

7. When finished, remove 96 well PCR plate (slot 1), which contains RNA extracted from samples. Move to Station C. If samples will not go through station C immediately, plate should be sealed and stored at -20℃ for short term storage or -80 for long term storage.

Overview of automated protocol

  1. Add IPA to samples. Mix.
  2. Add mag beads to samples. Mix.
  3. Magnet on. Remove supernatant (RNA stays with beads).
  4. IPA wash: magnet off, add IPA, mix, magnet on, remove IPA.
  5. EtOH wash: magnet off, add 80% EtOH, mix, magnet on, remove EtOH (x4)
  6. Dry beads (blow air over beads for 35 mins using constant pipetting)
  7. Magnet off, add nuclease-free water, mix
  8. Magnet on, move supernatant to 96 well plate

Summary of changes to standard protocol for automation

  1. Removed DNase step. Our tests showed this to be unnecessary for our particular qPCR setup (which is RNA-based, i.e. DNA does not interfere). In fact, the extra step only degraded the RNA in the samples.
  2. All mixing/spinning/centrifuging/vortexing has been replaced by a mix function, which pipettes the liquid up and down at a variable rate. Mixing times have been shortened drastically.
  3. Time required for magbeads to congregate to the side of the well appears to be almost nil; i.e. upon magnet engaging, the magbeads move almost instantaneously to a cluster at the side. Congregation time reduced to 30 seconds, to allow for any remaining congregation.
  4. Beads dry without the aid of a hotplate. Air is pipetted over the beads while they dry; our tests indicate that 35 minutes appears sufficient

Automation Notes

  1. Removal of EtOH before drying step is critical to successful drying. Aspiration of supernatant directed to bottom of well for this step, and excess volume aspirated.
  2. To prevent dripping, extra air must be aspirated before moving a volume of liquid across sample wells, tip racks, or other reagents.
  3. Excess volume must be dispensed when removing liquid from tips in order to fully void all reagent.
  4. During mixing, uneven volumes may be found in the various tips within the multichannel. Excess volume must be aspirated before mixing, and dispensed after mixing, to fully void the tips of liquid.

Automaction Code

Visit for up to date protocols.

Station C: qPCR and Analysis

In Station C, we have designed an automated protocol for adding one-step RT-qPCR master mixes to the extracted RNA samples and plating each reaction into the final qPCR plate for RT- qPCR . We based our RT-qPCR reaction around the uniplex SARS-CoV-2 assay from the CDC14. We used the N1 and the RNaseP primer-probe mixes in our assay. As we use a uniplex assay, it is necessary to run two RT-qPCR reactions for each sample, one for the N1 target and one for the RNaseP target, which acts as an extraction and sampling control. Both reactions for each sample were conducted on the same plate, however, to conduct RT-qPCR reactions for a full 92 samples, requires setting up two PCR plates for each sample plate.


Below equipment is for a single container lab  Station A capable of running 2400 tests in 24h.
Link to equipment inventory list.

  • 1 OpenTrons OT2
  • 2 Real-Time qPCR machines (96-well plate)
  • 1 Temperature module (OpenTrons)
  • 1 P300 8-channel (OpenTrons)
  • 1 P20 (8-channel ok) (OpenTrons)
  • 1 Undercounter Freezer (manual defrost!)
  • 1 PC (Windows) for qPCR machines
  • 1 Manual Pipette 200ul 8-channel
  • 1 Manual Pipette 20ul (8-channel ok)
  • 1 Goggles
  • 1 Labcoat


Test method: RT-qPCR

Below will provide an overview of the used RT-qPCR testing method from swab to result.
The Testing system consists of 3 main processes:

Sample preparation

Workflow Overview

There are 4 main ingredients that go into a swab system

  • Sterile Swab – the picture shows an NSP or nasopharyngeal swab which is has been shown
    to be the best system for extracting a sample from a patient for downstream applications.
  • Sterile tube – To store the sterile transport media before the sample is taken.
  • Sterile transport Media- A liquid solution to preserve the sample. (A dry sample can be used
    and PHE wales have demonstrated this). Of the liquid media there are couple of liquids
    a. Viral transport Media from a manufacturer- such as Virocult
    b. Opensource Viral Media- See appendix for recipe of such a solution
    c. GITC(Guanidine thiocyanate solution)- A solution alternative to chloroform for RNA
    extraction.(link to original paper)
  • Autoclave bag – to package everything together in a sterile environment

Sourcing these kits | There are two main options

  • Manufacturers- such as Copan
  • Opensource – It is possible to source these kits using 3D printing for the swabs and generic
    falcon tubes.

Taking a sample workflow

  • Rip open packaged swab system retain:
    a. Container containing VTM and NSP SWAB (Dispose of the autoclave bag safely!)
  • Take the NSP swab and insert through nostril. See this video for a better understanding.
    Please note that the video does not contain a NSP that requires snapping.
  • Place swab into container and snap, ensuring the swab remains in VTM
  • Firmly attach lid to swab
  • Follow PHE guidance on packing swabs for transport (insert document here).

Outputs | There will be two outputs

  • A Closed tube with VTM and Swab(send for testing)
  • A handle (DISPOSE of safely)

RNA extraction