Guidelines

1. Use the below Google Sheet to source parts for each of the eVOLVER subsystems (click on the tabs for different parts of the eVOLVER)

2. Parts may not be available due to backorder, discontinuation, or regional differences

3. If you are able to find an alternative vendor for a part, please post to the forum so it can be added to the spreadsheet

4. Should you not be able to order a part, post to the forum

eVOLVER Specific Parts

Vial Caps

Other "Consumables"

See Consumables tab in spreadsheet above.

Recommended to buy several vial racks as well for ease of handling vials.

Relevant Forum Posts

Check out the forum!

What are the different platforms used for?

Forum

For questions, discussion, and knowledge building

Should users find a problem, they should characterize it and submit it to group discussion on the forum

GitHub

For tracking and distributing code and hardware development.

Not for discussion

Wiki - Documentation

Basic information about the eVOLVER (about the project, contributing, where to buy)

Guides

In-depth hardware / software documentation

How to Edit the Wiki

Publications

Background, final data and experiments

Extensive documentation in the supplements, meant for reference

eVOLVER Community Code of Conduct

Useful Links

Software Repositories

Hardware Repositories

Buying

If you wish to purchase an eVOLVER, email account@fynchbio.com for a quote.

Connecting to eVOLVER

Currently eVOLVER requires a dedicated computer connected to it to run experiments. We recommend getting a Mac, which are easier to use and more reliable.

The computer needed for running experiments does not have to be new or powerful.

Upgrades

Variants

Khalil Lab

The Khalil lab uses eVOLVER to enable fundamental eco-evolutionary studies of cellular systems, and to evolve proteins with new and biomedically-useful functions at scale.

• Brandon Wong

• Zachary Heins

• Daniel Hart

• Ezira Wolle

• Nathaniel Borders

Alumni

• Chris Mancuso

Congrats on getting your eVOLVER and supporting our open-sourced community. We rely on support from you guys to help sustain this ecosystem so we can let everyone do the experiments that they want to! In this tutorial, we are going to teach you how to put this system together:

If you ordered the standard kit, this is the tutorial for you! The kit contains the following major components:

• Vial Platform

• 16 Smart Sleeves

• 2x Pump Arrays (total 32 pumps)

• Fluidic Box to for control of 32 Pumps (can expand to 48 w/ additional hardware)

1) First Unboxing the System

There are several components with protective foam. Insure all the components were delivered intact.

16 Smart Sleeves and a Box of Accessories

Pump Array and Fluidics Box

Vial Platform

2) Set up power and ethernet for the vial platform

The Vial Platform is the unit on which all the Smart Sleeves will be screwed onto. This system contains the eVOLVER Motherboard, Raspberry Pi, and Power Modules that control the Smart Sleeves. Parts used to setup the Vial Platform should be labeled with a black dot.

Contents of the Vial Platform Components Bag:

3) Plug in and screw on Smart Sleeves

If the system turns ON properly, turn off the eVOLVER and start fastening the Smart Sleeves. Use the 4-40 screws to fasten down each Smart Sleeve (2 screws/ sleeve). Vial 0 is in the front left (closest to the touch screen) with vials 1, 2, and 3 on the right of vial 0, in that order. Vial 15 is in the back right (farthest from the touch screen).

4) Setup pump rack

The Fluidics Platform is a separate module than the Vial Platform, typically is placed next to the Vial Platform. For this next part, we are setting up a pump rack that goes on top of the Fluidics Platform.

First we want to set up the components for the pump rack. The parts for this is located in the Fluidic Box Components Bag (Red Dot).

We want to fasten (w/ M6 screws) in aluminum extrusion into 3D printed part for the front pieces first. The front piece are the ones with the longer feet. See below:

Next, insert back piece. Make sure the two feet are offset in the same direction. Do not screw the screws in yet. We will screw those in when we mount everything together with the acrylic piece.

Repeat for the other side.

Next, line up the acrylic piece to the back holes and fasten together with M6 screws.

Screw in 10-32 screws to the feet of the mounts to fasten the pump rack on top of the Fluidics Box. Repeat to fasten all four mounts

5) Plug in Power + Communications to Fluidics Platform

Next, let’s setup the fluidics box. The fluidic box requires several connections:

• Two Power Entry Cables (one for 12V Power, other for 5V)

• Serial Communication Cable (To communicate with Raspberry Pi in Vial Platform)

Power the 5V power in the Fluidic Box from the USB port of the Raspberry Pi.

Connect Serial Communication Cable between the Vial Platform and the Fluidics Platform.

6) Attach Pump Arrays to the Fluidics Box

The fluidic box is meant to be flexible, to power whatever fluidic components you want. The standard setup assumes the following setup (back of fluidics box):

Attach ribbon cables to the appropriate slots on the PCB the pumps are mounted on.

Top down view of ribbon cable connections.

Finally, attach the tubing to the appropriate pumps.

7) Label Smart Sleeves with Appropriate Label

Page under construction!

Link to tutorial on forum for DPU installation

How to add multiple evolvers on the same computer

Can go through building router, but contact institution IT (tell them it's RPi)

Mac Installation

1. Download the evolver-electron-X.X.X.dmg file, where X denotes the version number in the latest release. The latest release should be the first one you see at the top of the page.

2. When the download finishes, open the file (it should be in your Downloads directory) and drag the eVOLVER app into your Applications directory.

3. Open the eVOLVER app from the Applications directory.

4. Using an editor of your choice (TextEdit, Nano, Vim, TextWrangler, etc.) open the file config.json located at /Users/<your username>/Library/Application Support/eVOLVER/config.json

5. Add a line in this file between the braces: "dpu-env": "/Users/<your username>/Document/dpu/venv"

6. Save the file and you're done!

Windows Installation

1. Download the evolver-electron-X.X.X.exe file, where X denotes the version number in the latest release. The latest release should be the first one you see at the top of the page.

2. Open the .exe file.

3. Using an editor of your choice open the file config.json located at C:\Users\<Your-UserName>\AppData\Roaming\eVOLVER\

4. Add a line in this file between the braces: "dpu-env": "C:\\Users\\<Your-UserName>\\Desktop\\dpu\\venv"

Note the escaped \ , be sure to put double \\ throughout the path.

5. Save the file and you're done!

Linux Installation

1. Download the evolver-electron-X.X.X.AppImage or evolver-electron-X.X.X.deb file, where X.X.X denotes the version number in the latest release. The latest release should be the first one you see at the top of the page.

2. If downloading evolver-electron-X.X.X.deb, open your Downloads directory and right-click evolver-electron-X.X.X.deb to open the file with Software Install .

3. A new window should open, allowing you to install the eVOLVER GUI. Click Install and follow the onscreen instructions to install the eVOLVER GUI.

4. Open the eVOLVER app by either clicking Show Applications from the home screen or through the command line. To open via command line, open Terminal and enter to access installed applications:

cd ~/usr/bin

Run the eVOLVER GUI by entering:

evolver

5. Using an editor of your choice, open the file ~/.config/eVOLVER/config.json .

6. Add a line in this file between the braces to denote the location of the installed DPU virtual environment:

"dpu-env": <path_to_dpu>

7. Save the file and you're done!

You can run eVOLVER experiment culture routines directly on the command line, or through the Electron graphical interface. This page documents how to install the DPU for use on the command line.

(Mac only) Homebrew and openssl/sqlite installation

In Terminal, install homebrew:

/bin/bash -c "$(curl -fsSL https://raw.githubusercontent.com/Homebrew/install/HEAD/install.sh)"

Afterwards, run the two following commands:

brew install openssl

brew install sqlite

Install Python 3.9

Python 3.9 is required for dpu operation

Mac:

brew install python@3.9

Windows

DPU Installation

2. Click Releases on the right side of the page

3. Click the Source Code link (zip) to download the latest release.

4. Move the downloaded .zip file (which should be in your Downloads directory for your browser) to the location where you want your experiments to run. Documents or Desktop are good locations. This is also where your data will be saved.

5. Unzip the file.

• If you are using a Mac and have installed homebrew (described above) you can run the following command: brew install poetry

• Windows Powershell:

(Invoke-WebRequest -Uri https://install.python-poetry.org -UseBasicParsing).Content | py -

8. Navigate to the DPU directory you just downloaded and unzipped:

• Use this command: cd <path_to_dpu>

• Replace the <path_to_dpu> with the location you put your dpu

Mac:

python3.9 -m venv venv

source venv/bin/activate

Windows PowerShell:

py -3.9 -m venv venv

venv\Scripts\Activate.ps1

10. Use poetry to install all necessary dependencies.

poetry install

If you can't find poetry on Windows "poetry : The term 'poetry' is not recognized as the name of a cmdlet" and it is installed, use the following command (replace <Your_User_Name> with your user name)

C:\Users\<Your_User_Name>\AppData\Roaming\Python\Scripts\poetry.exe install

Page under construction

Log in to router and make it have a static IP

otherwise IP will change each time it's power cycled

Then when connecting via ssh it will warn you and you need to change the file

You can then change the name, MAC address, and IP of the evolver in the evolver-config.json file

Router IP: 192.168.1.1

Login: admin

pw: password

About

In order to appropriately interpret data from and actuate physical elements connected to the eVOLVER framework, a relationship must be established between the electrical signals generated by attached sensors and the physical phenomenon they are measuring. This is done by manually measuring values on a controlled set of data via some gold-standard assay or measurement device and comparing these data to the eVOLVER generated data. By doing this across a range of different values, a function can be fit to this relationship to allow eVOLVER to interpret data accurately.

Procedure

Calibrations should proceed in the following order:

Vial Cap Upgrades

Benefits / Options

1. Better seal to vial - enables emergency efflux (see below) away from vial if too much is pumped in

2. More ports available

3. Reusable connectors - can use luer connectors to connect to eVOLVER tubing

Emergency Efflux

Protect your eVOLVER hardware with a sealed vial and additional efflux line dedicated to preventing vial overflows.

Bubbling

Many organisms require additional gas exchange outside of what eVOLVER stirring can provide. Others require custom gas mixes for growth. In-vial bubblers provide this.

Stop Stir While Taking OD

This is a measure that is necessary if you are bubbling, but can improve OD in general by decreasing noise.

You will need to calibrate with stir off. Swirl the vials when moving standards to make sure cells do not settle.

1. Fill a large beaker with water and submerge all of the pump lines (inputs and outputs of the pumps), ensuring all of the ends are below the surface of the water.

2. Run the pumps on the Setup page to completely fill the lines with water (about 20s)

3. Place 16 vials into a rack and place the output ends of the pumps to be calibrated into the vials.

4. Select the eVOLVER on the top of the GUI home page to be calibrated

5. Navigate to the Pump Calibration page

6. Enter a name for the pump calibration

7. Select the pump configuration for the eVOLVER. In a standard setup, IN1 are the media in pumps, and E refers to the efflux pumps. If you have another pump array (for a second media input source), it is typically going to be in IN2. Click Start Pump Calibration. The standard pumps are the FAST pumps (~1 mL/s, black pump heads). If you have the slow flow rate pumps (~ 1 ml/m, pink pump heads), select the SLOW radio button for that array.

8. Select all of the vials either by clicking the button on the bottom right or clicking and dragging across the vials on the selector. Click PUMP IN (or whatever the pump array you are calibrating) to run the pumps for 10s if the FAST radio button was selected or 100s if the SLOW radio button was selected.

9. After the pumps finish pumping, measure how much water was pumped by pouring out the water from each vial into a 25 mL graduated cylinder (or something similar). Enter this number into the box for each vial. The flow rate will appear on the vial selector on the right side of the screen.

10. Repeat this process for each pump array you have selected for calibration.

11. Click the pen button to submit and save the calibration. Then click Exit after the calibration is logged.

12. Verify the calibration appears on the setup page.

About

Why are temperature calibrations important?

Each microorganism has its own culture temperature requirement. Temperature may affect the successful cultivation and growth characteristics of the culture. By using a simple thermistor and heating resistor pair, we can track and change the culture temperature over time.

Why is calibration necessary?

Readings sent from the Raspberry Pi to the computer/server are the raw thermistor values measured by the Arduino. To make sense of the readings in terms of values we understand, we need a calibration file that converts this to degrees (celsius).

Do I need to redo temperature calibration if I switch out a thermistor/resistor pair?

Yes. Variability exists across the individual thermistor and heating resistor elements, necessitating a new temperature calibration anytime one of these parts are replaced. This is why the calibration curve is unique to each smart sleeve.

Materials

• 16 eVOLVER vials filled with 15mL water
• Thermometer with probes

  • Do not use a thermocouple-based thermometer, as the magnetic field from the stir components can interfere with measurements
• eVOLVER Electron GUI

  • This tutorial is for the newest version of the eVOLVER Electron App (Release 2.0.1), which enables both data visualization between each temperature calibration step (RT -> +10C -> +20C), as well as GUI-enabled calibration (without use of the command line).

  • For older versions of the Electron GUI (Release v1.2.1 and earlier), additional notes are provided at the bottom to successfully complete the calibration - the process remains largely the same, apart from the use of the command line to complete the calibration.

• eVOLVER

Methods

• Turn on the eVOLVER 5V power supply. Wait 5 seconds. Turn on the eVOLVER 12V power supply.

• Open up the eVOLVER Electron GUI app, and ensure you are connected to the correct eVOLVER you wish to calibrate in the upper-right hand corner of the "Home Screen". The circle should be solid green, indicating that the eVOLVER is connected.

• Tap "Setup" in the GUI home screen, select all vials by tapping "Select All" in the bottom right corner. The vial circles should highlight orange. Tap the "Next ->" button in the upper right-hand box (which includes all the control parameters) until you reach the "Temperature" tab. Drag the slider all the way down to "20C". Tap the button, and the "Executed Commands" box in the lower right should indicate that a temp command was sent to all vials.

• Place all the eVOLVER vials, pre-filled with 15mL of water, into the Smart Sleeves.

• Go back to the "Home Screen", click "Calibrations" and then "Temp" in the eVOLVER Electron App.

• Name the calibration, hit enter, and then click "Start Temperature Calibration". A prompt will read "Heaters turned off. Let equilibrate, then enter values."

• Room Temperature: Wait ~30 minutes for the temperature of the water to equilibrate to ambient room temperature (and heat generated by the eVOLVER).

• Once equilibrated, measure the temperature of each vial with the thermometer probe and log the temperature measurements in Celsius in the Electron GUI.

• Wait 60-120 minutes for the temperature of the water to equilibrate. Use the system outlined in the section "Temperature Equilibrium" to establish when the temperature has equilibrated.

• Wait ~2-3hrs, or leave overnight.

• Calibrate: An edit icon button should appear once all 3 measurements have been made. Click on the button, and a graph of the linear calibration curves for all thermistors across the smart sleeves should appear. Please confirm that the curves look linear. If everything looks good, please exit out of the graph (press the backwards arrow twice), and voila! You have completed your Temperature Calibration! Please be sure to change the temperature calibration file to the most recent in the eVOLVER set-up menu (click on the temperature calibration button).

About

The GUI will run calibrations automatically upon completion of the calibration protocols. However, you can still manually run a calibration if you would like to change calibration settings. Can be useful if your GUI calibration fails for some reason.

Requirements

1. You have run calibration and logged raw values already

Commands

List raw calibration files on the eVOLVER

Mac

For Windows, use py instead of python for all commands.

Calibrate Temperature

Calibrate OD

OD135

OD90 (Check to ensure mode is configured properly)

3D FIT (Check to ensure mode is configured properly)

Checklist

Before starting experiment prep:

1. Finished Getting Started Page

2. Calibrations

   1. Temperature (must be first, affects OD cals)

   2. OD

   3. Pump calibrations

3. Should things not be as expected

Sample Experimental Workflow

Day 0:

Make sure vials / pumps are behaving as expected

• Are all vials stirring, getting up to temperature, all pumps pumping, etc?

• Especially check that efflux is running long enough to remove enough culture from vials

Start overnight cultures

Day 1:

Inoculate cultures into vials

Monitor experiment for correct behavior (over several hours)

• Are the cells growing?

• Is the experiment diluting as expected?

• Are all pumps working?

Day 2 - Onwards:

Monitor media levels - make sure that there is more than enough media to make it to the next time you check the experiment

Monitor biofilming on the vials - this can cause increased OD over time

Relevant Forum Posts

Why

When we start an experiment, the script asks us whether we want to calibrate to blank.

Therefore, we take the first OD measurement as a zero and subtract that out from every subsequent measurement.

Details

OD blank is only applied to the calculated OD from calibration, not the raw OD data. The blank is applied for each vial, it just subtracts it out from subsequent measurements for that vial

1. Place waste lines into carboy

2. We use a small stand to bring the carboy closer to the eVOLVER

3. Have an extra carboy ready to swap in when the first gets full

4. (Optional) You can fill 10% of the carboy with bleach

   1. Or bleach after depending on whether you hate the smell bleach or your culture more

6. (Optional) Drill a hole through the carboy cap to make the opening to the carboy smaller (less smelly)

7. (Optional) Parafilm the opening to the carboy to make it less smelly

Materials

Media Requirements

A primary consideration when running eVOLVER experiments is predicting the media consumption rate, which is typically orders of magnitude higher than in typical batch experiments. The required media for continuous culture can be estimated using the following equation: 𝑀𝑒𝑑𝑖𝑎 𝑅𝑒𝑞𝑢𝑖𝑟𝑒𝑑 [𝑚𝐿] = 𝐶𝑢𝑙𝑡𝑢𝑟𝑒 𝑉𝑜𝑙𝑢𝑚𝑒 [𝑚𝐿] / 𝐴𝑣𝑔. 𝐷𝑜𝑢𝑏𝑙𝑖𝑛𝑔 𝑇𝑖𝑚𝑒 [ℎ] ∗ 𝑁𝑢𝑚𝑏𝑒𝑟 𝑜𝑓𝐶𝑢𝑙𝑡𝑢𝑟𝑒𝑠 ∗ 𝐷𝑢𝑟𝑎𝑡𝑖𝑜𝑛 [ℎ] (1) It should be noted that additional media (~20mL) is needed during device setup to flush media input lines.

Protocol

1. Before preparing media, steam clean the carboy by filling with ~2L of water and autoclaving on the liquids cycle for at least 45 minutes. Be sure to keep the lid loose to prevent the carboy from collapsing. Pour out the water.

2. If you are able to prepare a concentrate of your media, you can do so as a 10x 2L solution (or whatever makes sense for your media) in a beaker with a stir bar. It is also possible to mix everything directly in the carboy. Add your media components in small batches and try to mix as you go.

3. Fill the carboy up to 20L - volume of media components that will be supplied after the autoclave step. For example, if you need your media to be 2% glucose, you should fill the carboy to 18L and then add 1L of filter sterilized 40% glucose after autoclaving.

4. Place a male Luer cap on the outlet of the carboy lid. Place a filter on the inlet, and cover it with foil. Loosen the entire lid.

5. Place the carboy into an autoclave bin, then place into an autoclave. Autoclave on the liquids cycle for 2 hours.

6. Take the carboy out of the autoclave.

7. If media components need to be added after autoclave (glucose, antibiotics, supplements, etc.) place the carboy on a cart and move next to a bench flame. Sterilize the area thoroughly with ethanol before quickly unscrewing the lid to make the additions. Mix by placing the carboy on a cart and moving the cart in a circular motion, or by placing on the floor, tilting ~30 degrees, and moving the top in a circular motion.

8. You can place the autoclave on the floor next to your experiment. Be sure to keep the lid loose to prevent negative pressure buildup in the carboy which could prevent media from going into the culture vessels.

9. After the carboy is empty, rinse thoroughly with water, being sure to run water through all of the tubing. Steam clean as described in step 1, then pour out the remaining liquid.

Relevant Forum Posts

Running experiments with Dropbox/Google Drive

If you would like your data to be automatically backed up on Dropbox or Google Drive, first install the desktop applications. This will enable any computer with Dropbox/Google Drive and access to the directory to monitor the experiment using the eVOLVER GUI in real-time, while providing all the benefits of using these services (automatic backup of data, ease of data sharing, etc).

Setup

Select the eVOLVER you wish to run an experiment on in the dropdown on the top right, then click the SETUP button.

On left side of the screen, ensure the correct calibrations are selected for Temp, OD, and the pumps. Click the box to bring up a list of available calibrations for each parameter.

Once you have selected the calibrations, you can set the initial stir and temperatures, and sterilize the fluidics as described in the JoVE paper.

Experiment Manager

The JoVE paper is out of date with regards to using the GUI for running and managing an experiment, from 3) Configuring eVOLVER Software and Programming Algorithmic Culture Routines on.

Open the experiment manager on the Home screen of the GUI. You should see something that looks like this if this is the first time you have used the application:

In the top center the Expt Directory is listed out. You can click the pen icon to change this location to be anywhere you like.

If you have Dropbox or Google Drive installed, you can select a location on the cloud to save your experiments. This will enable you to view your data from any computer that has permissions and the Dropbox/Google Drive application installed!

You do not need to make a new folder called experiments as shown below - the GUI will automatically create this directory.

Click the New Experiment button to open a prompt to name your experiment. Type a name then click Submit.

Click the Pen button on the row of the new experiment to bring up with experiment editor page.

Experiment Editor

By default, the Turbidostat editor will appear when you come to the editor page.

1. The name of the experiment. You can click the pen icon to change the experiment name.

2. The parameter selector. The name of the currently selected parameter is listed at the top. You can click the -/+ buttons to change the value (or use the slider), and then click the box in the middle to apply this value to the selected vials.

3. Experiment control panel. From left to right: Start/Stop experiment, View Data, Save Files (Only for File Editor), Copy Experiment, View Files in File browser, Delete experiment.

4. eVOLVER Selector. Select the eVOLVER to run the experiment on.

5. Experiment type. Default will bring up the Turbidostat editor. Chemostat and growth rate editors are also available. You can also bring up the file editor to directly modify the python files within the GUI.

6. Vial selector - select the vials and then apply desired experimental parameters from the buttons on the parameter selector. Values display the currently set parameters.

7. Vial selector control buttons - flip the orientation on the display or select all of the vials.

8. Reset - goes back to default. Save Experiment - saves the parameters for all vials.

The chemostat and growth rate modules function similarly to the turbidostat one.

Clicking File Editor in the experiment type dropdown will allow you to directly modify the python files.

You can select the file you would like to edit on the left, edit it on the right, and save by clicking the Save icon on the bottom left. For most cases you will not need to use this! If you change the files, you will not be able to use the t-stat/c-stat/growth rate modules, and you will need to modify the EVOLVER_IP, OPERATION_MODE, and other variables throughout the code.

You can also click the folder icon on the bottom left to view the files in a file browser and edit them using an editor of your choice.

Starting the experiment

When you're ready to start the experiment, click the Play button either on the Experiment Manager page or the Experiment Editor page. Then you can click the graphs icon to go watch your experiment running in real-time.

When you navigate to the graphs page, the graphs will look fairly empty to start. After a minute or so, data will start to appear as it is collected. You can also click the VIEW LOGS button to see how the experiment is running.

On the left, you can modify how the graphs are displayed by clicking the appropriate radio buttons. You can also look close up at an individual vial and use the data slider at the bottom of the graph to zoom in on an area of interest.

When you click ALL to go back to seeing all vials, the range set on a single vial will be applied to all vials. You can click the radio buttons to set it back to the latest data.

To end the experiment, click the STOP square button on the device running the experiment.

Code Breakdown and Explanation of Flow Rates

• OD_data: The OD_data variable saved from the transformed values from the input_data variable. The data is transformed via calibration scripts called from eVOLVER.py.

• start_OD: This variable determines if you want the culture to be above a certain OD before starting dilutions

• start_time: This variable determines when the dilutions should start. If set to 0, the dilutions start immediately

• OD_values_to_average: The script will average N recorded OD values before it triggers the dilution via the start_OD variable.

• chemostat_vials: Modify this script to to turn off vials (e.g. pumps). There is a known bug on the rate definition that might not turn off the pumps there (see below).

The peristaltic pumps are fixed flow rates, does that mean I have to use different pumps for each flow rate setting?

No, with the eVOLVER setup, we approximate continuous perfusion by frequent small boluses of media. By doing this, a faster pump can achieve tunable, slower net flow rates. In our experience, this compromise does not significantly impact the culture conditions of common laboratory microbes.

Here is an example calculation one would make:

• The culture volume is 30 mL/ vial.

• The typical pumps eVOLVER comes with have a fixed flow rate of 0.5 - 1.0 mL/s. We typically recommend a minimum bolus of 0.5 mL per dilution, meaning 60 boluses would turnover the culture once.

  • FynchBio also sells slower pumps that flow at ~1 mL/ min

• If we want our culture to double every 3 hours, the pumps will fire every 3 minutes (180 min/ 60 boluses). In a chemostat, nutrient is limiting thus growth rate == dilution rate.

• A doubling time of 3hr has a growth rate of 0.231/hr via the following equation:

Below, we set the script such that the cultures will all have a 3 hour doubling:

If one wanted to set the V0-3 at a doubling time of 3 hours but the rest of the 14 vials with a doubling time of 6 hours, the rate_config would be set as the following:

NOTE (5/12/2023): There is a bug that does not turn OFF pumps if defined to 0 on rate_config variable.

Code for interfacing with eVOLVER GUI

Below is code that grabs values from the GUI and populates it in the code. If you aren't using the GUI, please ignore this portion of the code:

Rest of the Chemostat Configuration Settings

The following parameters initialize variables that will calculate how much time to turn the pumps ON to maintain the flow rate set in rate_config

• flow_rate: get the calibrated flow rates for each pump

• period_config: how often to pump (seconds)

• bolus_in_s: how long to pump during a given pump event (seconds)

Chemostat Pump Commands

When we set vials to a new flow rate in the chemostat function, we see the experiment script send a comma separated pump command. Each pump assigned to the a chemostat gets a command that looks like this:

1.18|144

Which is in the format of:

bolus_in_s|period_config

If the above pump was influx, its corresponding efflux pump would be 2.36|144 because we pump out double the time to avoid overflows.

Stopping Experiment and Bleaching

1. In the GUI electron app or command line, click to stop your experiment.

2. In large beakers, prepare 500mL of 10% bleach and 200 mL of 10% bleach.

3. Disconnect media bottles and place lines into 10% bleach solution

4. To use pumps to bleach the cultures, keep tubing connected, but move vials to a white eVOLVER rack. Prepare an additional 1L of 10% bleach and add to the media input beaker.

5. Run 10% bleach through vials until final concentration in the vials reaches ~10% bleach (~60 seconds for 25mL volume). Disconnect influx and efflux lines from vials, set vials aside for 30 mins.

6. To manually clean vials, disconnect influx and efflux lines and skip steps 39 and 40

7. Submerge the disconnected influx and efflux lines in the 200mL bleach beaker, then run all pumps for 20 seconds to fill all lines with 10% bleach. Let sit for no less than 30 minutes, no more than 48 hours.

8. Remove media input lines from bleach, and run air though the lines to remove bleach. If not planning on using the device again right away, run diH2O through all lines for 20 seconds to remove residual bleach from the lines. Dried bleach can form salt crystals and clog the lines. If this happens, break up the crystals manually using pliers to squeeze the line.

9. Turn off the 12V power, then the 5V power.

10. Proceed to vial & bottle cleaning protocol.

Clean Vials After Bleaching Cultures

1. Prepare a beaker ~50% full of 10% bleach

2. Remove vial lids and place in 10% bleach solution

3. Using the magnetic stir bar grabber, remove the stir bars from each vial, place in the 10% bleach solution

4. Let vial caps and stir bars sit for 30 minutes-overnight in the 10% bleach solution

5. Meanwhile, dump bleached culture down the drain, rinse with diH2O

6. Use a test tube brush to scrub inside of vial, particularly important if you grew bacterial cultures

7. Place bleached/scrubbed vials right-side-up in the vial racks to indicate they have not yet been dishwashed

8. Once vial lids/stir bars have been soaked for sufficient time, carefully pour out the 10% bleach solution, and replace with diH2O

9. Let vial lids and stir bars soak for another 30 mins-overnight.

10. Once vial lids/stir bars have been soaked for sufficient time, carefully pour out the diH2O.

11. Run diH2O through influx and efflux straws for 2-3 seconds at the sink to remove any residual bleach

12. Place stir bars in the stir bar tray.

13. Once 60+ vials have accumulated, vials should be dishwashed.

14. Place vials open side up in 3 white eVOLVER racks.

15. Take one rack, cover the vials using the wire mesh from the bottom rack of the dishwasher, then flip upside down. Repeat for the other two racks.

16. Add any other glassware or eVOLVER bottles to the top rack as desired, then run on cycle 3.

17. Store vials upside down in vial boxes to indicate that they have been dishwashed.

Ways you might accidentally change the OD reading

1. Moving vials once experiment has started

2. Altering

Overview

Reasons you might use command line experiment start:

1. Doing a custom experiment

2. Unable to install GUI or getting errors you aren't able to fix easily (post on the forum first)

Protocol

1. In a command line window, enter the dpu virtual environment.

   1. Navigate to the DPU directory

      • Use this command: cd <path_to_dpu>

   2. Enter the command:

      • On Mac OS:

        • source dpu-env/bin/activate

      • On Windows PowerShell:

        • dpu-env\Scripts\Activate.ps1

2. Navigate to your experiment folder

   1. This should be in dpu/experiment

3. Copy the template folder and rename it for each experiment

   1. Navigate into your new experiment folder

4. Alter settings in custom_script.py

   1. EVOLVER_PORT = set correct IP of the eVOLVER

   2. Set initial temp and stir settings
5. Start the experiment using the command:

   1. python3 eVOLVER.py -i <your_evolver's_IP_address>

   2. python3 eVOLVER.py -i 192.168.1.9

6. 1. Temperature should be fully equilibrated before this

   2. DO NOT add cells before blanking, whatever OD value the cells + media have will be subtracted from future measurements

7. Stop the experiment using the keyboard shortcut control+C

   1. One control+C pauses the experiment from recording data

   2. Two control+C stops the experiment

8. To change parameters mid-experiment or restart from blank data files:

   1. Stop the experiment fully

   2. Change parameters in custom_script.py

   3. Restart the experiment

      1. Continue to keep your data

         1. Overwrite to make new blank data files and throw away your old experiment. NOTE: you will lose your OD blank and will need to make a new one (a hassle if you have already inoculated with cells)

9. You can run experiments on more than one eVOLVER at a time

   1. Create a new experiment folder (different custom_script.py) and change IP

   2. Follow this procedure in an additional command line window

Ending an Experiment

Overview

While large carboys (autoclavable plastic jugs >10L) are useful for long-term experiments or particularly fast-growing bugs, the best typical media storage is standard Pyrex/Corning bottles, which range from 100 mL to 10 L. The 1-2L bottles are the most convenient size for media prep, short experiments, and experiments with many media types, so we'll be sharing the process for those bottles here.

Parts:

1 ft of 1/8" ID silicone tubing 2 Female luer-lock connector with 1/8" barb, polypropylene 1 Male luer-lock connector with 1/8" barb, polypropylene 1 Male luer cap Tools: Scissors/razor Drill gun 3/16" drill bit

Step-by-step

First, drill a hole directly down through the cap. Be sure to have something safe to drill into on the other side. If having trouble with the 3/16 bit catching on the plastic, drill a pilot hole with a smaller bit.

Next, push one of the barbed female luer connectors through the top of the cap, so the luer side is facing up.

Next, cut two pieces of tubing. One is for the cap above, and can be as short as 3". Put the two remaining luer adapters (one male, one female) on this piece. This flexible piece makes it a little easier to connect lines to bottles. The other should be at least 9" so that it makes it from the barb on the cap all the way to the bottom of the bottle (for 1L bottles). Connect the longer piece of tubing to the barb poking through the bottom of the lid.

The long straw should reach the bottom of the bottle and bend slightly. If it’s too long it will be annoying to screw the bottle cap on, and might get stuck against the wall. If it's too short, it won’t be able to reach media at the bottom of the bottle. Your bottles may vary: for 2L bottles, you need a 13" straw.

Rinse out the straw with diH2O, and add a male luer cap to the top of the straw before autoclaving, and you’re good to go!

You can also make splitters with the same 1/8" tubing and corresponding luer-barb connectors and barbed T-connectors, allowing you to split one media bottle to multiple input lines. We usually sterilize the splitters using 10% bleach and ethanol (by connecting them before sterilizing all the lines when beginning an eVOLVER experiment), but everything shown here is autoclavable as well.

Background

eVOLVER hardware is primarily divided between the Smart Sleeves and Motherboard. The Motherboard is where all the sensitive control hardware is, whereas the Smart Sleeve is nice modular way of integrating all the sensors to control your culture conditions.

Goal

The goal of this tutorial is to show how one can build an eVOLVER Smart Sleeve using commonly found tools and easily customizable parts.

Time/Cost

That all sounds great, but I have never soldered or 3D printed before, how do I start?

How can I modify or change the design?

Does this tutorial include how to build the Motherboard?

No. This tutorial only provides instructions on how to build a Smart Sleeve.

Guide

1. Order all of the necessary parts and tools.

List of supplies needed ( displayed in photograph above ):

1. Laser Cut 1/4" Acrylic Base

2. 2 pieces of laser cut 1/8" Acrylic Fan Spacers

3. 5/64" Hex Key

4. IR LED

5. IR Photodiode

6. Thermistor

7. 2x Socket head screw

8. 12 V DC Computer Fan with Magnets Glued

9. Vial Board

10. Lab Tape

11. 3D Printed Part

12. Aluminum Tube

13. Soldering Iron + Solder

14. 2x Stainless Steel Screws, 2.5" 4-40 threading (Not Shown)

2. Familiarize yourself with how the Vial Board, 3D printed part, and aluminum tube should be assembled.

Take notice if all the components line up properly:

• Tapped 2-56 holes on the Aluminum Tube lines up with holes on heating resistors (2 black rectangular components soldered on vial board)

• The heating resistors should sit on two rectangular holes on the 3D printed part

• Green screw terminals line up with the LED holes on the 3D printed part

Take note of where the two holes labeled “thermistor” are on the PCB.

3. Tape thermistor on the aluminum tube near where the two vias were located on the PCB.

When taping the thermistor, ensure the writing on the thermistor is facing away from the aluminum tube. Tape the thermistor parallel to the length of the tube about .5 cm away from the bottom of the tube. The leads should be exposed when slotted into the 3D printed part.

4. Slot the aluminum tube into the 3D printed part. Verify that the holes for the LEDs on both the aluminum tube and 3D printed part are properly aligned.

Make sure 3D printed part is slotted tightly. Add additional pieces of tape if is loose. Avoid damaging the thermistor by twisting or rotating the parts.

5. Align the PCB over the 3D printed part, with the two thermistor leads threading through the vias on the PCB.

Also verify that the two small 2-56 holes on the aluminum sleeve align with the holes on the heating resistor.

6. Use the 2x Socket head screws to fasten the heating resistors to the aluminum sleeve.

Aligning the hole with the heating resistor makes screwing the socket cap in much easier. Thermal paste can also be applied before this step to form better contact between the heaters and aluminum tube.

7. Place the 1/8" laser cut acrylic pieces onto of the fan to act as spacers between the magnets and glass vessel.

8. Use the two 4-40 stainless steel screws to fasten the assembled 3D printed part and the computer fan with spacers. Fasten both screws onto the 1/4" acrylic piece.

9. Coil the wires from the computer fan around the Smart Sleeve and connect to appropriate ports on the screw terminal labeled “fan”.

The red wire should go into PWR and the black wire should go into the GND ports.

10. Attach the LED (clear packaging) and photodiode (dark packaging) for OD measurement to the appropriately labeled slots.

Both the photodiode and LED should have a short and a long lead. This corresponds to the polarity of components. Make sure for both components, the longer lead is connected to PWR . If this is done improperly, you will not get a reliable signal.

11. Bend the LED and diode into the corresponding slots.

12. Finally, solder the thermistor onto the PCB.

That’s it! You’re done.

Turn on the eVOLVER 5V power supply by flipping the associated switch. IMPORTANT: Wait for 5 seconds before turning on the 12V power supply.

Ensure the correct eVOLVER is selected by consulting the upper right hand corner of the GUI. It will display the eVOLVER name (default is evolver-darwin), followed by the IP address in parentheses. This will be the IP address you assigned it when setting up the router connecting the DPU to the vial platform.

In the PARAMETER box, ensure the correct calibrations are selected for Temp, OD, and the pumps. Click the box for each to bring up a list of available calibrations for that parameter.

Select all vials by tapping Select All in the bottom right corner of the screen. The vial circles on the right side of the display should become orange.

You can also click and drag to select as many vials as you want.

Tap the NEXT → on the box that says parameter to get to the Stir tab. Tap the - button or drag the slider to a Stir of 8. Press the Stir: 8 button to alter the stirring of all vials.

Tap the NEXT → on the box that says parameter to get to theTemperature tab. Tap the - button or drag the slider until the middle button says 37.0 °C and tap the button. The EXECUTED COMMANDS box should state that a stir and then temp command was sent to all vials.

Pumps

About Server Logs

We also provide a utility monitoring and restart script, evolvercron and server_monitor.sh. To use, add the crontab job line in evolvercron into your cron installation.

View Server Logs

1. Connect to your server via the following command (replace <eVOLVER_IP> with your eVOLVER's IP)

   1. ssh pi@<eVOLVER_IP>

2. Enter the eVOLVER server's password

[Option 1] View the server log live

1. Enter the supervisor environment via the following command:

   1. sudo supervisorctl

2. Use the following command to follow the eVOLVER server log:

   1. tail -f evolver

3. Ctrl+C to exit

[Option 2] View the whole server log file

You can either view there or move it to your computer for more analysis

Restart the eVOLVER Server

2. Enter the following command:

   1. restart evolver

3. If you quickly follow the server via tail -f evolver you should see it restart

Overview

Reasons you might want to do this:

Guide

1. In a terminal, navigate to the folder in the server files containing the conf.yml file you want to change to

   1. For example if you want to change to the stir pause conf.yml:

      1. cd <your-directory>/evolver/alternate_conf_files/stir_pause_for_OD_reads

   2. For example if you want to change to the default conf.yml:

      1. cd <your-directory>/evolver/

2. Copy over your new conf.yml file (overwriting the old one)

   1. Input the following command:

      1. scp conf.yml pi@<your-IP-address>:/home/pi/evolver/evolver/

   2. Input your eVOLVER server's password

3. Restart your eVOLVER server

   1. Input the following command (replaced with your eVOLVER's IP)

      1. ssh pi@<your-IP-address>

   2. Input your eVOLVER server's password

   3. Input the following commands to restart the server

      1. sudo supervisorctl

      2. restart evolver

4. Check the server log files to see that it is cycling properly

   1. While still in supervisorctl, input:

      1. tail -f evolver

Using Raspberry Pi Imager

3. Plug an 8GB micro-SD for the RPi into an adapter, then into a the computer.

4. Click the Choose OS button, then scroll to the bottom and click Use Custom.

5. In the popup at the dropdown at the bottom of the window, select All files(*.*). Then select the image file downloaded in Step 1.

6. Click the Choose Storage button and select the SD Card.

7. Click the Write button. A notification might appear warning about erasing all existing data on the card. Click Yes. If it asks for your computer password afterwards, enter it. It might take a while to finish (~40m).

Using command line utilities (Mac/UNIX only)

2. Plug the SD card into the computer.

4. Run the following command:

diskutil list

Identify the SD card name from this (it should be obvious). Next run:

diskutil unmountdisk <diskname>

where <diskname> is the name you identified from the previous command - it should look like

/dev/disk3/ or something similar.

5. Use the dd command to copy the image onto the SD card:

sudo dd bs=4m if=<diskimage.img> of=<diskname>

Backing Up Raspberry Pi (Make a Custom Image)

Writing an image to save your full Raspberry Pi configuration.

Useful to save time when reimaging your Raspberry Pi if you want to go back to your custom Python code and calibrations as quickly as possible.

Page under construction

Protocol

Control Layer (1/4" acrylic)

Cut outlines first

• 0.1pt

• 3 copies

Keep full stock in place + outline on computer

• Acts as a frame for the device

• Remove board from stock with tweezers

Add adhesive

Put adhesive covered outline back in place on stock

On the computer

• Align control vector cuts (ports) to the outlines

• Delete outlines

• Print (3 copies for 1/4")

Keep board in place

Rastering control channel

• Undo deletion of outline, delete previous vector cuts

• Align raster outline to old outline + delete outline

• Load control_channel 2 in advanced settings

  • 1200 dpi

  • Deep to get through adhesive

• 1 copy => print

Remove board + stock from laser cutter

Flow Layer (1/8" acrylic)

Cut adhesive sheet

• Copy/paste in design with scooped out region around bridge

• Cut adhesive out first

• This way we don't have to move the acrylic stock later to put adhesive on the laser cutter

• Check adhesive stickiness

  • • Remove corner from adhesive

    • One side will stay attached to adhesive, one will remove cleanly

• Put adhesive sheet on laser cutter, with side that easily removes down

• Focus to adhesive

• Vector cut Full speed + 80 - 90% power

Cutting acrylic

• Delete scoops, we just want the outlines

• Print vector cut for outline

• Refocus laser cutter to 1/8th inch + cut

Plasma clean acrylic

Put adhesive on

Rastering acrylic

• Place device back in stock on laser cutter

• Copy/paste raster of flow route

• Remove outline

• Raster cut

• While rastering, start tapping control board

Assembly

Tap control board

• 2 turns one way, little turn the other way

• Use a countersink to remove burrs around the holes

  • A couple of turns

  • Both sides of board

  • Allows silicone to sit flush + barbs to insert easily

• Blow out small bits of acrylic from threading using air before plasma treating

Remove backing from all acrylic (helps with alignment later)

Silicone

• Sam from Densmore lab bought a big roll

• Some in drawer under cutting station

• Cut larger size than device

• Peal one side of silicone

• Plasma treat 30 seconds

Apply silicone to control board

• Align and press onto control board

• Take off plastic from other side of silicone

• Wrap in paper towel

• Put between aluminum plates in vice

• Turn tight + one small more turn

• Wait 10 minutes

Punch holes in silicone

• Use biopsy punch (green on shelf above 3D printers) to make holes silicone for flow

• Don't need to do the control

• Put pressure and turn

Clean silicone + adhere to flow layer

• Use piece of packing tape to remove dust

• Plasma clean silicone 30 seconds

• Align flow and control

  • Take special care to align IPP flow and control

  • Easier to align flow layer on top

• Clamp full device in paper towel for 10 minutes

  • Avoid over tightening - this can overstretch the silicone layer

Put loctite on barbs + screw in to board

• Barbs are in Dan's drawer

Subprotocols

Laser Cutter

Different colors of line produce different depth / intensity of cut

• Use black for everything

Make all lines hairline outline

• Otherwise it won't cut!

Open up a new page

• Paste whatever you're cutting into this

• This will be the correct size of bed

Printing

USB Bullshit

• Open up printers / devices

• Have to reconnect USB multiple times for it to be recognized

1. Open up print

2. Select correct printer you figured out

3. Load the correct settings

4. Set the correct copy number

5. 1. For millifluidics this is 2 for vector cutting (we have the double line problem)

   2. 1 for raster probably

6. Press print

On the Laser Cutter

Make sure it's on / connected

Focus first using metal thing attached to laser

Run the job with lid open to double check it's doing what you hope

Run the job

Plasma Cleaning

Plasma treat surfaces (except adhesive)

• Take off one side of acrylic backing

• Vacuum knob arrow left

• Metal valve tight, then quarter turn loose

• Pump on + power on to remove air 1 minute

• Then switch on upper right knob to high

  • Acrylic = 1 minute treatment

  • Silicone = 30 seconds treatment

• Turn off pump and power

• Slowly open black knob and let vacuum fill

Adhesive

Adhesive is in a box above 3D printers

Cut adhesive to larger size than acrylic

• Remove adhesive backing and place face up on surface

• Remove acrylic from plasma cleaner and firmly press clean side onto adhesive

• Squeegee the bubbles out as much as you can (and away from where the channels will be)

• Cut excess adhesive off with razor blade

Page under construction

While batch culture techniques often utilize biosafety hoods or flame convection currents to keep workspaces sterile, these are rarely amenable for automated cell culture devices. Prevention of contamination in eVOLVER is achieved at three levels: 1) sterilization of media, culture vessels, and fluidic lines, 2) attention to sterile technique, and 3) physical and chemical barriers to contaminants. First, all media bottles and their adapters, and all components of the culture vessel (e.g. borosilicate glass vial, magnetic stir bar, cap with fluidic adapters) are designed to be autoclaved before each use. Fluidic lines on the device are sterilized before and after each experiment using bleach and ethanol (see Online Methods). Second, following sterilization, sterile technique should be practiced when attaching media lines to culture vials by working quickly, avoiding physical contact with the ends of fluidics lines, and taking care to spray gloves with ethanol. Finally, additional physical and chemical measures may be taken depending on the organism and experiment. For example, the sampling port may be covered by a sterile membrane for long-term culture. For slow-growing eukaryotic cultures, antibiotics can be added to the media to exclude bacterial contaminants. UV sterilization of components or surfaces is another preventative measure to consider.

Page under construction

Plug in 5V power supply and 5V fan

Changing Pressure Settings

• Unplug the SAMD21 from the pressure regulator before uploading code (otherwise there might not be enough power to the SAMD21 from the USB)

• Upload modified pressure code to SAMD21

Option 2: Add a 'pres' command to the evolver conf.yml and modify conf.yml

Fluidic Lines

Connect a 8-10 psi air input into the input port (only port on its own in 1.0 design)

Connect regulated line to both sensor (below) and controlled output (white, above) as shown below:

Set Up Solenoid Control

Solenoid Bank Diagram (facing holes)

• The solenoids should be normally open and connected to the 8 psi air source. ie Vacuum when closed

• Left = 8 psi

• Right = Vacuum

• Middle = 8 different solenoids, groups of 3 are IPP controls

Connect solenoid ribbon cable to slot 5 on the fluidics box

Turn on eVOLVER server and fluidics box

Check that the solenoid bank is correctly set

Run ipp_cal.py on the server:

• Use command:

  • python3 ipp_cal.py <frequency (hz)> <seconds to run>

  • For example: python3 ipp_cal.py 1 100

Set Up IPPs

• Hook IPPs up to a pressurized input bottle for consistency

• Make sure that IPPs are operating as expected and producing the expected flow rate

Miscellaneous

Overview

Optical density (OD) is one of the most integral parts of continuous culture. While chemostats do not innately rely on OD, it is still a useful metric for your evolution. Meanwhile, turbidostats and growth curves heavily rely on OD to function.

However, OD is the most easily disrupted parts of eVOLVER hardware. This guide serves as an initial list of things to check when troubleshooting OD.

Most Common Problems

Variability in OD Sensing Range Between Vials

2. If you already have a version of the 3D printed tube holder with defined OD LED and photodiode distances:
5. Try different ADC resistor packs

   1. eVOLVERs come with a variety of these

   2. Ask on the forum for help

6. Check that your vial construction is not variable

Noisy OD Data

1. Stir bars jumping around during OD readings

   1. Decrease stir speed
2. You are near the edge of your calibrated OD range

Incorrect OD

ie eVOLVER OD is offset from the cuvette or plate reader measured value.

1. 1. Especially because you did not wait for temperatures of the vials to equilibriate

2. Poor quality OD calibration

   2. Different temperatures require different OD calibrations

No Change in OD

1. Photodiode or IR LED not working

   2. A failed photodiode will show a raw OD value close to 64,000

   3. A failed IR LED will most likely show a raw OD value much closer to your other vials

   4. Check that they have the long (positive) lead in the correct positive terminal

2. ADC board or motherboard broken

   1. Can happen after vial overflow

Other Considerations

Pre-Calibration Considerations

• od_led value in conf.yml file (PWM value)

• • Use the splash guard that comes with your eVOLVER

  • You may need to recalibrate if you calibrated with ambient light

Experiment Considerations

• Photodiode Temperature - make sure temperature is equilibrated before calibration and before start of experiment

• eVOLVER recently turned on - causes rapid change in values

• Needles or other structures coming below 10mL line (close to OD PD etc)

• Vial volume - a volume of 10mL will totally change OD compared to 30

• Stirring with low vial volume - this will give noisy readings as the funnel created by stirring fluctuates and is more and less reflective

Non-standard changes

• Vial holder reflectiveness - ie white vs black vs metallic

Overview

If a parameter is not working properly, often the first place to check for issues is the server

Guide

1. Monitor the server log file
2. Any time you are troubleshooting the server, make sure to save your calibrations.json file in a secure location before going further

3. If the log file is not updating (via evolver tail -f)

   1. If you have manually edited the conf.yml file, most likely you have made an error somewhere there

      1. This could be formatting or a typo

      2. 2. ONLY copy the conf.yml to home/pi/evolver/evolver/

         3. Otherwise you will overwrite your calibration files

      3. Restart the server

         1. Stop following the server using ctrl+C

         2. Type restart evolver

         3. evolver tail -f

      4. The server log should now be updating each cycle

Relevant Forum Posts

Relevant Forum Posts

Vial overflow is one of the most common problems to plague eVOLVER experiments. Overflow events can cause damage to internal components such as the PCB motherboard, sensor actuator (SA) boards, power supplies, etc. If you experience overflow during an experiment, it is important to immediately stop the eVOLVER and evaluate potential causes of overflow, as well as any damage that the overflow may have caused.

If you catch an overflow that happened recently (i.e. an overflow that happened overnight), the best thing you can do after stopping the experiment is turn off and unplug the eVOLVER, determine the extent of overflow spilling, and wash any components that have media on them. There are no components on the vial board or motherboard that retain significant charge (i.e. no batteries or large capacitors), so they are safe to wash gently in soap and water. Once clean, they should be dried quickly with a paper towel.

The main reason for electronics failure after an overflow is current shorting through media (most microbial media is salty and conductive), or connections and copper corroding from media that has been left to dry. If you can clean off all the media immediately after an overflow, you will have a much better chance of saving electrical components, and saving yourself many headaches down the road.

If you did not catch an overflow quickly, or a large amount of media spilled, follow the workflow below.

Troubleshooting Workflow for Overflow

Inspecting internal components

Overflowed media can enter into the vial platform through the ports that connect the Smart Vials to the motherboard. This can cause damage to the internal circuitry, which is should first be evaluated by visually inspecting the motherboard, SA boards, power supplies, and Raspberry Pi. Damage to any of these components can look like burned or melted spots or loose/broken connections. If you see visible damage to any of these parts, they will need to be fully replaced. See pages below for specific instructions on how to replace these various parts.

Old evolver pumps (solid black plastic head)

Relevant Forum Posts

Stir Bar Jumping

Additional tip: if you see strange raw OD values after swapping out LEDs or photodiodes, it's possible you have the positive and negative ends switched.

Description

As IPP devices are sensitive to changes in pressure at valves and in connected media bottles, we developed an 8-channel pressure regulator that can be used to regulate these pressures through the eVOLVER framework. The device consists of sets of two proportional valves that can limit air flow from a high-pressure source and a vent at atmospheric pressure. By connecting an electronic pressure gauge to the output of this valve configuration, it is possible to implement proportional-integral-derivative (PID) control over the valves in order to set the output pressure to any desired level between the input and atmospheric pressure. We validated the functionality of this device by regulating pressure at 1.5 psi over 24 hours, and compared the performance of our device with that of a fixed, manually set regulator (PARKER-WATTS R25-02A) connected to the benchtop air supply (Supplementary Figure 3). The average pressure with PID control was 1.498 psi with an RMS error of 0.0086 psi, while the fixed regulator had an average pressure of 1.706 psi with an RMS error of 0.2220 psi. Large pressure deviations (>0.5 psi) that can affect the performance of the devices were observed with the fixed regulator, but were successfully eliminated with our automated pressure regulator scheme. We further characterized the effects of pressure changes at various locations in the system in order to optimize performance of the IPP devices for the course of a PACE experiment (Supplementary Figure 3).

We have monitored the lifetime of eVOLVER components since the invention of the device (~3 years). As in most systems, mechanical parts have the highest possibility of wear and thus need replacement most frequently. Likewise, in eVOLVER, the peristaltic pumps used have a lifetime of 6 months of typical use during continuous culture. Specifically, the silicone tubing within the head of the pump is frequently compressed when actuating peristaltic pumps and will tear over time. The head can easily and inexpensively be replaced ($4/ pump). The computer fan used for stirring has robustly operated continuously for > 3 years, whereas the magnetic stir bars are rated for a limited number of autoclave cycles and need eventual replacement. All other components (e.g. heaters, thermistors, LEDs, diodes, PCB, power sources) have been stably operating for > 3 years.

Standard components

The standard eVOLVER peristaltic pumps (since ~2020) sourced by Fynch Bio are selected specifically to eVOLVER use. Previous pumps used with eVOLVER during its early development tended to wear out quickly and have variable flow rates, primarily due to the mechanism for rotating the rollers against the tubing to drive pump flow. The motor shaft directly turned the rollers through friction, which led to slippage and abrasion over time. The current pumps do not have drirect contact between the motor shaft and rollers, instead driving roller rotation through a gear reduction system.

This gear system drives a roller casing with fixed roller axles contained in the pump head. In this way, the pump action does not rely on friction and thus much more accurate and durable than previous pumps.

Troubleshooting and repair

Because the current pump style has been introduced so recently, very few of them have failed from wear and tear. Based on our experience with other pump styles, the most likely failure mode will be collapse or tear of the tubing inside the pump head, or shearing of the tubing on the barb outside the head. This can be solved by replacing the pump head (contact Fynch Bio) or replacing the tubing.

The induction motor inside the pump is very unlikely to fail through normal use. If there is a liquid spill on the pump array and some pumps are not spinning, the most likely cause is corroded contacts with the pump array PCB. While the pumps themselves will be fine, they should be removed and replaced to a new PCB, ordered from Fynch or PCBWay. This will require removing solder, which can be tricky.

Swapping and modifying

The pump heads can be rotated 90 degrees in any direction, should your bench setup require changing the direction your pumps face.

The low-flow pumps available through Fynch use an additional gear reduction box to reduce the speed of pump action by a factor of 45. This is useful for addition of inducers, antibiotics, or other media feeds that require a small and precise bolus.

Plenty of other peristaltic pumps are suitable for eVOLVER, so long as they operate at 12V.

The flow rate of eVOLVER pumps (and any pump with an induction motor) is somewhat modifiable through PWM modulation. This effectively reduces the voltage going to the pump, which will slow its rotation, down to the point where the coils can't generate enough torque to consistently turn the shaft. This happens about halfway through the PWM range (around 2000), where you can get the pumps to run at about 1/3 speed. This is rarely necessary for experiments, so it has not been optimized, but is a potential feature for users to explore.

From eVOLVER Forum

Overview

Diagnosis

If you have determined that the server and Arduinos are both functioning properly, however, the link between them seems to be broken in some or all ways, then the last likely culprit is the motherboard.

In the example below, overflow events during an experiment caused media to leak through the Smart Vial ribbon cable ports into the vial platform, which houses all of the eVOLVER circuitry. This resulted in visible damage to the motherboard, which can be seen below.

Replacement Protocol

Unfasten Motherboard

The motherboard is fastened to the eVOLVER platform and other internal components in several ways (numbers correspond to image below):

1. Ribbon cables connecting the motherboard to the Smart Sleeves (not shown)

2. Screws in gold holes

3. 6 screws that fasten wiring from the power supplies to the motherboard

4. Cable connecting motherboard to Raspberry Pi (fastened to base of platform)

5. Cable connecting motherboard to display screen (fastened to lid of platform, obstructed by ribbon cable in image)

Remove Motherboard

1. Unplug the cables connecting the motherboard to the Raspberry Pi and display screen.

2. Remove the screws in the gold holes using a Phillips screwdriver.

3. Remove the screws holding the wiring in place using a flathead screwdriver.

4. Close the eVOLVER chassis lid half way, making sure to hold the motherboard with one hand. Unplug the ribbon cables that connect the Smart Sleeves to the motherboard, starting from the last row of vials and moving towards the first row of vials.

5. Maneuver the motherboard around the 3D printed part housing the display screen.

Transfer Parts to New Motherboard

The PWM boards, ADC boards, SAMD21 mini Arduino boards and jumpers need to be transferred to the new motherboard.

Remove the ADC boards, PWM boards, and SAMD21 mini breakout boards from their slots

1. Using an Integrated Circuit (IC) puller, as seen in the image below.

   1. Pull the IC puller perpendicular to the motherboard to prevent bending or breaking the SA boards’ pins.

2. Or put a screwdriver in the hollow under the board and SLOWLY work it back and forth the pry the board out.

Make sure you pull the pins out as straight as possible because it will be challenging to get the board back in if they are bent.

1. If pins are bent in the process of removing the boards, they can be straightened using tweezers. Avoid bending the pins too much; excess bending can cause the pins to snap off.

2. If pins are snapped off during this process:

   2. Alternatively, the pins can be soldered back into place onto the board, but this is much trickier to do.

   3. The board should be replaced with a new board if other options do not work

Optional: If your motherboard includes a variable resistance board, proceed to steps 3 and 4. If not, skip to step 5.

1. Remove the variable resistance board from the motherboard. The ADC boards on this board can be removed first, or the variable resistance board and ADC boards can be removed as a single part.

2. Remove the headers under the variable resistor board and transfer to the new motherboard. These can be removed with pliers.

2. On the side of the resistor array, there is a description of the part and a black dot, seen in the image below. This black dot indicates the ground pin of the resistor array, and should go into the slot towards the top of the motherboard.

1. Remove the jumpers to the old motherboard by pulling them straight up. Transfer the jumpers to the new motherboard, recreating the same configuration as shown below.

1. After checking for damage, transfer the variable resistance board (if applicable) and SA boards to the new motherboard (it is easiest to move these boards after transferring the resistor arrays and jumpers). Be sure to align the boards’ pins with the slots on the motherboard before pushing the boards into place.

Adding the SA parts to their slots on the motherboard can be quite tricky, especially if their pins were bent during the removal process. Some tips for this step:

• Inspect boards and use tweezers to carefully straighten pins that have been bent out of place. Bending the pins back and forth too much can cause them to break off the board, so this should be kept to a minimum.

• If pins break off an SA board, they can be soldered back into place, or the entire board can be swapped out.

• Place the SA board over its slots and check for pin alignment. Try to wedge the bent pins into place first, before pressing the board into place.

• Make sure ALL PINS enter their respective slots before pressing the SA board into place.

• The SA boards can withstand some force when being pushed into the motherboard. Press the SA boards into their slots until the boards lay flush with the slots on the motherboard.

1. Unscrew the 3D printed corners from the old motherboard and fasten them to the new motherboard. These parts act as spacers to prevent the backside of the motherboard from touching the eVOLVER platform.

Prepare New Motherboard and Plug in to eVOLVER

1. Use electrical tape to cover all exposed elements on the back of the motherboard. There are exposed pins on both the top and bottom of the motherboard.

The new motherboard is now ready to be fastened to the vial platform!

1. Align the motherboard correctly by lining up the ribbon cable ports with the openings in the platform.

2. Fasten the motherboard to the platform using the screws and a Phillips screwdriver.

3. Fasten the wiring from the power supplies to the motherboard. The new motherboard will have screws in place for where the wires should go, so those should be removed and then used to fasten the wires to the motherboard.

4. Plug in the cables to the Raspberry Pi and display screen.

5. Plug in the Smart Vial ribbon cables to the motherboard from the top of the eVOLVER.

Power Up eVOLVER

You are now ready to power up your eVOLVER!

The first time you turn on the eVOLVER after replacing the motherboard, we recommend opening the platform and keeping an eye out for a few things:

• Be sure to turn on the 5V power supply first, wait 10 seconds, then turn on the 12V power supply.

• If you notice smoke or the smell of something burning, immediately turn off both power supplies. If this occurs, inspect the motherboard and the SA boards for melting or burn spots.

• The red light on the Controls Raspberry Pi should turn on steady, and the green light should blink on and off.

• The red and blue lights on the SAMD21 mini breakout boards should turn on steady.

What's in the box

The fluidics box can be opened by removing the 8 phillips head screws on the sides. Inside the box is the fluidics board (with a SAMD21 and three PWM boards) with RS485 and 5V connections for the SAMD21, and a 12V power supply beneath the board. Should the need arise to update the arduino code on the SAMD21, you'll need to open the box to get to it.

The connections to the pumps are numbered according to their address, i.e. cable 1 connects to pumps 1-8, cable 2 connects to pumps 9-16, etc.

Troubleshooting and repair

Not many things can go wrong inside the box, but we have dealt with a situation where the 5V power connection was faulty and led to malfunction. This part can be replaced by opening the box and replacing the faulty connector.

Modifications

Adding another pump array

The existing pump pox can control up to 16 additional pumps (or any other 12V valve, actuator, etc) with no modifications. Typically this would mean using connections 5 and 6 on the back of the box, which correspond to indices 33-48.

Modifying fluidics code

Daisy-chaining multiple boxes

Two RS485 ports (the telephone jack-looking ports) are available on the back of the fluidics box so that multiple boxes can be linked together on the same RS485 "line". All SAMD21 microcontrollers are receiving the same serial data from the Raspberry Pi, and only responding to commands when the appropriate address for that SAMD21 is in the command, like "temp" or "stir". Additional fluidics boxes can have the same "pump" address with higher pump indices (i.e. pumps 49-96 for a second box), so long as you change the number of expected pump values on the conf.yml file. You will also need to modify the SAMD21 arduino code accordingly. You can also have a different address for the other fluidics box, by adding another parameter in the conf.yml and modifying the arduino code accordingly.

Materials

• 3D printed vial cap with ports for nylon tubing

  • • Service = SLS; material = nylon PA-12

    • Get "vapor smoothed" for better sealing

• Mixing tray (here a small weighboat)

• Disposable applicator (here a wooden stir stick; pipette tip also works)

Design

1. Choose the minimum number of ports necessary

   1. You will need to block up ports that are unused if you want to do emergency efflux

   2. Therefore, choose the cap with the minimum number of ports that you need

   3. For example, counting ports for PACE

      1. Lagoon: 1 vial-to-vial + 1 efflux + 2 inducers + 1 emergency efflux = 5 ports

      2. Cell reservoir: 1 media influx + 1 efflux + 1 vial-to-vial + 1 emergency efflux = 4 ports
2. Efflux straw sets the volume of the culture

3. Increase influx accuracy

   1. Important for low volume influx like inducers

   2. Angled influx tubing should touch the inside of the glass vial to create a stream of liquid rather than drops.

   3. It should also be well above the efflux line to avoid contamination.

4. [Optional] Use different colors of tubing to denote different ports (see above)

Construction Protocol

0. Mark fluid lines on a glass eVOLVER vial (see below image)

1. This will be used to set the efflux tube height, which sets the volume in the eVOLVER vial.

2. Tape an eVOLVER vial

3. Using a serological pipette, measure water into the vial in increments of 5 mL

1. Cut Nylon Tubing to Various Lengths Needed

1. Cut the necessary components of the hard semi-clear white nylon tubing (1/8" OD). Refer to the table below for lengths.

The table below has the necessary components for the construction of one cap.

1. Cut the necessary components of the flexible nylon tubing. Refer to the table below for lengths.

The table below has the necessary components for the construction of one cap.

2. Assemble Cap

Steps

1. You may need to drill out the ports in the 3D printed vial cap if powder is blocking them.

2. Insert the hard semi-clear white nylon tubing (1/8" OD) into each corresponding port in the cap as per the image below.

1. Screw the cap into the glass vial with measuring tape on the side and ensure that:

• The 2 influx tubes ARE touching the edge of the vial. They should be staggered such that one is higher than the other.

• [Bubbler Only] Bubbler tube is at the 15mL line on the vial.

• [Bubbler Only] Bubbler IS NOT touching the edges of the vial and is facing the sampling port (rotate and adjust as necessary to ensure this).

1. Follow the epoxy directions to mix up a small amount of epoxy in a disposable dish. Use even pressure to get even amounts of resin and hardener. Use a toothpick or other similarly sized applicator to put the minimum amount of epoxy around the outside area of the cap where the ports and the hard nylon tubing connect, ensuring to spread the epoxy in between any gaps.

2. Set assembled caps upright (with a vial rack or spare vials) so that epoxy doesn't drip into the sampling port as it cures.

3. Set cap on its side to cure overnight.

4. Next day, add the appropriate color flexible nylon tubing (3/32" ID) to each hard nylon tubing (1/8" OD) as per the table above.

   1. [Bubbler Only] Connect the flexible nylon tubing (3/32" ID) bubbler connector component onto the inside end of the gas bubbler hard tubing. Allow the tubing to reach halfway through the connector tubing.

   1. [Bubbler Only] Connect the constructed gas bubbler onto the other side of the connector tubing. The end of the gas bubbler hard tubing that is inserted in the cap and the gas bubbler should connect as per image below.

Standard components

Tubing

Silicone is the tubing of choice for eVOLVER because of its flexibility, durability, cost, and resistance to beach and ethanol. It is also autoclavable, although 30 minutes of 10% bleach sterilization is sufficient to completely sterilize the lines.

Connectors

The barb side of the connector is rated for the ID of tubing it is mean to be used with, i.e. 1/16" barb for 1/16" ID tubing, 1/8" barb for 1/8" ID tubing, etc. Barbs are designed to get a tight seal around their widest diameter, with the max pressure and resistance to slippage depending on the softness of the tubing and style of barb (40 psi for standard eVOLVER connectors).

Luer connectors work by pressing a "male" plug into a "female" socket, both tapered at 6%. Because of the smooth surfaces and large contact area, this creates an effective seal without tools, held in place by friction. Syringes and medical equipment commonly make use of luer connections.

To hold luer connections in place more securely, luer-lok connectors use a screw-like "skirt" to prevent luer connectors from simply being pulled apart. eVOLVER tubing connections are luer-lok, allowing for easy but relatively secure connections between most components.

However, luer-lok connections have some common failure modes. Because the threads of the skirt are so coarse, it only takes a 1/4 turn to break the seal at the plug-socket interface, leading to a leak (if positive pressure) or air bubble source (if negative pressure). Loose luer connections easily become disconnected, leading to spills (the major source of electronics failure) and of course, failed experiments. One should always ensure luer connections are snug (hand tight), and tubing free from torsion that might unscrew the luer-lok.

Additionally, because the male connector comes in direct contact with fluid, it becomes your most likely contamination source. Be careful not to bump the tips of the male connectors against contaminated surfaces when setting up your experiment. Doing a final wash with 70% ethanol after bleach sterilizing can prevent some contamination from this route, depending on your media and organism.

Swapping and modifying

Any soft tubing (durometer 40-70A) works well with the barbed connectors. Vinyl (PVC) or tygon tubing might be a suitable alternative for decreased permeability. Opaque tubing may make sense for light-sensitive media or antibiotic feeds that have extended time exposed to light in the tubing due to low flow rates, at the cost of not being able to observe media in your line. Remember that 1/16" ID tubing contains about 0.75 mL per foot.

Materials

• 3D printed (SLS, nylon PA-12) vial cap

  • Get these "vapor smoothed" for better sealing

  • Get white nylon PA12 (the default black will leach dye into your media)

  • Other default settings are fine

• • Buy at least 4 x number of caps + extra

  • Nylon required to survive autoclaving (expansion differential between normally used polypropylene and nylon 3D printed vial causes epoxy to separate)

Construction Protocol

1. [Optional] Mark fluid lines on a glass eVOLVER vial (see below image)

   1. This will be used to set the efflux tube height, which sets the volume in the eVOLVER vial.

   2. Tape an eVOLVER vial

   3. Using a serological pipette, measure water into the vial in increments of 5 mL
2. Using a razor, trim nylon tubing so that it reaches the volume you have set

   1. Nylon tubing will straighten in the autoclave, so try and straighten it out as much as you can

   2. Cut to 53 mm for 20mL

   3. Otherwise, determine the correct length using graduated eVOLVER vial from step 1

      1. Cut the tubing to a longer length than you need

      2. Pushing it into the vial cap until you feel a little resistance (easy to accidentally go further, but there is a stopping point)

      3. Screw the cap on all the way

      4. Estimate where to trim

      5. Check in vial cap again

      6. Measure length when you have this correct - use this for the rest of your vials (or use cut length as a guide for cutting the rest)

   4. Cut all tubing for the caps you are making

1. Use gloves when working with epoxy

2. Put epoxy into mixing tray and mix

   1. You don't need much because epoxy will likely set before you finish all vials

1. Apply epoxy to barb of luer connector

   1. Be generous, but avoid the opening in the barb

   2. If you do get epoxy in the opening, blow it out or pick it out with a paperclip

1. Press the connector into a port in the top of the cap, and twist a few times to spread epoxy

1. Repeat steps 4 and 5 for the other ports

   1. When epoxy starts to solidify, mix more (in a new tray)

2. Wait for at least an hour for epoxy to set somewhat

3. Epoxy one side of the outside of efflux tube (avoiding the hole)

4. Press the tube into left port of cap (when sampling port is facing you) (see below)

   1. Only press until you feel a slight resistance

   2. Twist once or twice to spread epoxy

1. Repeat steps 8 and 9 for all caps

2. Allow epoxy to set 24 hours before using

Description

The programmable Smart Sleeve is the foundational unit on which the eVOLVER is built (Fig. 2 and Supplementary Fig. 1). The Smart Sleeve is comprised of all the sensors and actuators required to control the culture conditions inside a 40 mL borosilicate glass vial. At the core is an aluminum sleeve, which surrounds the vial and is used to control temperature via two resistive heaters and a thermistor integrated within. Near the base of the vial sits a 3D printed part that houses and aligns the optical density LED and photodiode. Below that sits a fan motor equipped with magnets to rotate a stir bar within the vial. The Smart Sleeve represents one of the most easily customized features of the eVOLVER: by changing which sensors and actuators are used and their layout, the user may develop culture vessels that fit their experimental needs. For a detailed description of the sensors and actuators used to control stirring, temperature, and optical density in Smart Sleeves featured in this study, as well as strategies for modifying the Smart Sleeve to fit experimental needs, refer to Supplementary Note 4. Liquid handling is also controlled at the level of the individual culture vessel, yet these components are housed in a separate fluidic module, described in Supplementary Note 5.

The sensors and actuators on each sleeve are integrated in a small printed circuit board (PCB), termed the Vial Board (VB). We designed the VB such that we can easily solder electrical connections and efficiently manage/package wiring from the sensors. The VB is a very simple PCB, containing only a few resistors, and is straightforward to redesign and inexpensive to manufacture, if needed. The simplicity in the VB leads to robustness in the system. For example, any accidental overflow and spillage from the vials (e.g. from clogged fluid lines or user error) should minimally impact the rest of the system, as critical components are located at the Motherboard rather than the sleeve itself. Ribbon cables provide a modular way to connect the integrated Smart Sleeves to the Motherboard.

The VB is designed to rest atop a 3D printed piece, which houses optical density and temperature components (see Supplementary Note 4). The printed part can be fabricated with Nature Biotechnology: doi:10.1038/nbt.4151 5 any commercial or DIY 3D printer, readily available at almost any university or hacker space, and customized to the requirements of the user. For example, if a user wanted to change the mode of optical density detection between scattering and absorption, they could redesign the 3D printed part housing the LED-diode pair such that it would have the correct offset angle for the desired mode of measurement.

Vial

Vial Holder

Vial Sleeve

Vial Cap

Vial Board

Version D

Pin Map

Differences to Version C bolded

1. Fan

2. Unused; linked to fan for future fan tachometer

3. Heating resistor

4. Thermistor

5. OD IR LED

6. OD 135 photodiode

7. OD 90 photodiode / Spare A

8. Spare B

Version C

Pin Map

Differences to Version D bolded

1. Fan

2. Unused; linked to fan for future fan tachometer

3. Heating resistor

4. Thermistor

5. OD IR LED

6. OD 90 photodiode

7. OD 135 photodiode / Spare A

8. Spare B