Protocol to study human gut bacterial communities and rhythmicity ex vivo using a chemostat system

Summary

Chemostat systems can be used to cultivate complex intestinal microbial communities ex vivo. Here, we present a protocol to transfer bacteria from human fecal material into chemostat systems as well as settings to simulate infant or adult colonic conditions. We describe the experimental setup, media design, donor selection, 16S rRNA amplicon sequencing, and circadian analysis of bacterial abundance. This protocol enables the investigation of changes in microbial community composition and bacteria-derived metabolites upon exposure to different dietary components.

For complete details on the use and execution of this protocol, please refer to Heppner et al.¹

Graphical abstract

Highlights

• •Details of chemostat setup for continuous ex vivo cultivation of intestinal microbiota
• •Characterization of nutrient-microbiota interactions ex vivo
• •Versatile applications to simulate different intestinal conditions
• •Circadian analysis to determine rhythmic changes in microbiota composition and function

Publisher’s note: Undertaking any experimental protocol requires adherence to local institutional guidelines for laboratory safety and ethics.

Chemostat systems can be used to cultivate complex intestinal microbial communities ex vivo. Here, we present a protocol to transfer bacteria from human fecal material into chemostat systems as well as settings to simulate infant or adult colonic conditions. We describe the experimental setup, media design, donor selection, 16S rRNA amplicon sequencing, and circadian analysis of bacterial abundance. This protocol enables the investigation of changes in microbial community composition and bacteria-derived metabolites upon exposure to different dietary components.

Before you begin

Institutional permissions

Make sure that fecal sample collection follows ethical and institutional guidelines and is only performed after the required permissions have been granted. Data used in this protocol were obtained from intervention trials carried out in accordance with the Declaration of Helsinki and approved by the Ethics committee of the Technical University of Munich (study numbers 254/17S and 2022-466-S-SR).

Experimental design

Timing: weeks

• 1.
  The general experimental setup and timeline of ex vivo cultivations is described in Figure 1.

  • a.
    Set up and autoclave the cultivation system then check sterility at least 48 h by using Brain-Heart Infusion (BHI).
  • b.
    Exchange BHI for the first cultivation media during media change.

    Note: The timing of this phase is dependent on the applied speed of media influx (”feed rate”) and therefore variable.

  • c.
    During batch culture, inoculate the selected fecal material into the cultivation vessels twice, once at the beginning and once after 24 h of cultivation. This allows the bacterial community to establish.
  • d.
    To test multiple conditions (in form of cultivation media), divide the continuous culture into different phases.

    • i.
      A stable bacterial community is reached after 7 days of exposure to one cultivation media.
    • ii.
      For circadian analysis, rhythmicity sampling can be performed after establishment of stable communities following day 7. The rhythmicity sampling elongates the exposure to one cultivation media by 48 h.
    • iii.
      In case of testing multiple cultivation media, the media can be exchanged after day 7 of media 1 (or day 9 in case of rhythmicity sampling). Achievement of complete media exchange depends on the chosen retention time (see step 2). Rhythmicity sampling can be performed again after formation of stable communities with cultivation media 2 (following day 7).

      Note: The stability of bacterial communities decreases after ∼19 days of continuous culture (data not shown).

  • e.
    Perform the experiment in duplicates (i.e., two cultivation vessels with the same conditions).
• 2.Depending on the research question, use the following settings to replicate either adult or infant conditions in the cultivation system (see Table 1).
  Independent of the mimicked conditions, the following points apply for all experiments.

  • a.
    Media 1 should mimic the diet at the time point of sample collection (i.e., adult medium for adult samples).
  • b.
    The minimal operating volume for the cultivation systems is 400 mL, which corresponds to the average colon content of an adult person.⁷
  • c.
    To test the influence of certain ingredients, they need to be integrated into the cultivation media (see Section “experimental media design”).
  • d.
    Samples can be collected from the cultivation system at any time. Daily sampling should be performed at the same time of day. Sampling for circadian analysis should be performed at least every 4 h for 48 h. The sampling volume depends on the required volume for downstream analyses, such as:

    • i.
      Naive sample: gDNA isolation, metabolomics, plating
    • ii.
      20% glycerol: bacterial isolation, fecal microbiota transfer
    • iii.
      Additional analyses: RNA isolation, transcriptomics

      Note: Samples for metabolomic analyses can be processed and fractioned if necessary or snap frozen and stored at – 80°C until analyses.

  • e.
    In case of rhythmicity sampling or inclusion of photosensitive ingredients, consider covering the cultivation system (and media bottle and connected silicone hose) with aluminum foil. While doing so, ensure easy access to the vessels to verify media and gas influx.
  • f.
    The redox potential will be monitored but not controlled throughout the entire experiment.
• 3.Once the experimental design is determined, transfer all phases with the respective settings as well as the inoculation time points into eve® software (see Figure 2). Refer to “Plan and Run Multiple Batch” in User Guide eve® (Version 1.123, 2021 H2).

Note: eve® software is a manufacturer-provided software used for monitoring and recording of experimental parameters (i.e., pH, temperature, redox potential) throughout the experiment, which facilitates vessel comparison and troubleshooting.

Figure 1.

Setup and timeline of an ex vivo cultivation experiment

(A) Picture of an ex vivo cultivation system. Continuous arrows indicate flow directions. Dashed arrows point to the named accessories.

(B) Experimental timeline of an ex vivo cultivation experiment testing the influence of two different cultivation media on a microbial community. RS, Rhythmicity sampling for circadian analysis.

Table 1.

List of settings to mimic adult or infant conditions in the ex vivo cultivation systems

	Adult conditions	Infant conditions
Sample origin	Adult donor	Infant donor
Cultivation media	Based on adult diet2	Based on infant diet3
Retention time (feed rate into cultivation system)	36 h4 (11.1 mL/h)	12.5 h3 (32 mL/h)
pH	6.85	6.56

Figure 2.

Screenshot of phases of an exemplary experiment investigating the effect of two cultivation media in eve® software

The filled orange square marks the first inoculation. ∗pH set point depends on research question. ∗∗Feed rate during media change can be adjusted as necessary. ∗∗∗Feed rate during continuous culture depends on research question.

Experimental media design

Timing: multiple weeks to months

Note: The integration of target foods into the experimental media is important for the experiment’s outcomes and depends on the research question.

• 4.Calculate the daily media consumption (depending on retention time and working volume).

$$Dailymediaconsumption\left(mL\right)=\frac{Workingvolume\left(mL\right)}{Retentiontime\left(h\right)}\times 24h$$

• 5.
  Determine the amount of target food per L cultivation media as follows:

  • a.
    To mimic the human situation, calculate the daily intake of the target food (DI).
  • b.
    Refer to digestibility indices of individual ingredients (i.e., sugars) to determine the amount reaching the colon.

    Note: It is also possible to perform in vitro digestion of the target ingredients before integrating the residues of the digestion into the cultivation media.⁸

  • c.
    Calculate the amount per L cultivation media depending on the daily media consumption.
• 6.
  Exemplary calculation for cultivation media mimicking infant conditions according to Cinquin et al.³ (see Table 2).

  • a.
    For the amount of different carbohydrates, proteins and fats in the target food, refer to ingredient lists and nutritional values provided by the manufacturer (i.e., Töpfer GmbH, Germany).
  • b.
    Calculate the proportions of ingredients per compound class (carbohydrates, N_compounds, fats).

    $$Proportion(\%)=\frac{Amount\left(g/100g\right)}{Totalamountofcompoundclass}\times 100$$
  • c.
    Calculate the amount of ingredients per compound class in the daily intake of the target food.

    $$Amount\left(\frac{g}{gDI}\right)=\frac{Amount\left(g/100g\right)}{100}\times DI\left(g\right)$$
  • d.
    Calculate the amount of digested and undigested ingredient (g/ g DI) of each component by applying digestibility indices.

    $$Digestedingredient\left(\frac{g}{gDI}\right)=Amount\left(\frac{g}{gDI}\right)\times \frac{Digestibilityindex}{100}Undigestedingredient\left(\frac{g}{gDI}\right)=Amount\left(\frac{g}{gDI}\right)-digestedingredient\left(\frac{g}{gDI}\right)$$
  • e.
    Calculate the proportions of undigested ingredients per compound class (see step 6.d.).
  • f.
    Calculate the ratio of carbohydrates:N_compounds:fat of undigested ingredients.

    $$Ratio=\frac{Undigestedcarbohydrates\times 100}{Totalundigestedingredients}:\frac{Undigested{N}_{compounds}\times 100}{Totalundigestedingredients}:\frac{Undigestedfats\times 100}{Totalundigestedingredients}$$

    Note: To mimic digestion of fats in infants, the fat content was set to 10%.⁹

  • g.
    Set the total amount of undigested carbohydrates to 13 g/L media.² Calculate the amounts of individual carbohydrates by applying the proportions of undigested carbohydrates (see step 6.f.).
  • h.
    Calculate the total amount of N_compounds.

    $$TotalN\_compounds\left(\frac{g}{L}\right)=\frac{13\frac{g}{L}}{Undigestedcarbohydrates}\times undigestedN\_compounds$$
  • i.
    Calculate the amounts of individual N_compounds by applying the proportions of undigested N_compounds and subtracting 0.5 g/L for tryptone and peptone respectively.

    $$IndividualN\_compounds\left(\frac{g}{L}\right)=\left(TotalN\_compounds\left(\frac{g}{L}\right)-1\left(\frac{g}{L}\right)\right)\times Proportionofindividualcompound\left(\%\right)$$
  • j.
    Calculate the total amount of fat (see step 6.f.).

    Note: For experimental convenience and to decrease costs in case of the cultivation media mimicking the infant diet, fat was added in form of the target food (Organic Infant Milk Pre, Töpfer GmbH, Germany).

  • k.
    Calculate the amount of target food containing the total amount of fats.

    $$Amounttargetfood\left(\frac{g}{L}\right)=\frac{Totalfat\left(\frac{g}{L}\right)}{Fatcontentintargetfood(\%)}$$
  • l.
    Calculate the amounts of individual carbohydrates and N_compounds in the amount of target food (see step 6.k.) and subtract this from the values calculated before (see steps 6.g. and 6.h.).

    Note: Depending on the ingredients of the target food, the same applies to the remaining media components (i.e., salts).

• 7.Dissolve the ingredients of your media and sterilize it by autoclaving (121°C, 15 min).

Note: For heat-sensitive components, consider sterile filtration or autoclaving at 110°C for 15 min.

• 8.Check the solubility of all components before and after autoclaving. Adjust the media preparation if precipitates or a change in media color occur.

Note: It is possible to divide the compounds into multiple solutions that are sterilized separately and to combine them in a sterile environment.

• 9.
  Test the suitability of the designed media for the cultivation system and determine a suitable feed rate (%).

  • a.
    Connect the media bottle to a feed pump and the pump to an empty bottle using silicone hoses (Ø = 2 mm).
  • b.
    Set the feed rate to 100% and start the feed using the touch screen.
  • c.
    Monitor the media influx over at least 4 h. Regularly check for blockages in the silicone hoses and note the volume in both bottles.
  • d.
    Calculate the media influx (mL/h) for 100% feed rate. Calculate the feed rate necessary to apply the determined retention time (see Step 2).
  • e.
    Use the touch screen to set the feed rate to the calculated value and monitor media influx for at least 4 h. Verify that the media influx results in the determined retention time.

CRITICAL: High viscosity of the media increases the risk for blockages in the silicone hoses supplying media into the cultivation vessels.

• 10.
  The ingredients of exemplary media for adult and infant conditions are shown in Tables 3 and 4.

  • a.
    Prepare 1 bottle of media 1, 2 and 3 per cultivation system. Store at 4°C for up to 7 days.
  • b.
    Media 4 should be prepared freshly or stored at 4°C for up to 7 days.
  • c.
    Aliquots of media 5 can be stored at – 20°C for the duration of the cultivation experiment.
  • d.
    Combine media 1–5 in the bottle of media 1 in a sterile environment. Use sterile ddH₂O to fill up to final volume. Store combined media at 4°C until connection to cultivation system.

CRITICAL: To avoid degradation of media components, connect the combined media to the cultivation system no later than 7 days after autoclaving.

Table 2.

Exemplary calculation of the cultivation media mimicking the Organic Infant Milk Pre (Töpfer GmbH, Germany)

Compound class	Ingredient	g/100 g target food	Proportion (%)	Digestibility indices (%)	Digested ingredient (g/100 g target food)	Undigested ingredient (g/100 g target food)	Proportion undigested ingredient (%)	Ratio undigested ingredients (C:N:F)	Total amount (g/L)		Amount target ingredient (g) for 2.969 g fat	Individual ingredients (g) in 11.88 g target food	Add to media (g/L)
Carbohydrates	Lactose	60.000	99.932	983	58.8000	1.2000	99.932	44	13.000	12.991	N/A	7.126	5.865
	Inositol	0.041	0.068	9810	0.0402	0.0008	0.068			0.009	N/A	0.005	0.004
N_compounds	WPH	5.940	59.885	803	4.7520	1.1880	93.722	46	13.723	11.924	N/A	0.705	11.219
	Casein	3.960	39.923	983	3.8808	0.0792	6.248			0.795	N/A	0.470	0.325
	L-Carnitine	0.019	0.192	9811	0.0186	0.0004	0.030			0.004	N/A	0.002	0.002
Fats	N/A	25.000	N/A	N/A	N/A	N/A	N/A	109	2.969	N/A	11.88	N/A	N/A

Table 3.

Exemplary preparation of a cultivation media mimicking adult conditions

Media #	Ingredient	Amount (per L media)	Solved in	Sterilization method	Comments
1	NaCl	4.5 g	600 mL pre-heated ddH2O	Autoclave 121°C, 15 min	Remove lumps by filtering through sieve multiple times. Autoclave in bottle big enough for final volume (i.e., 2 L). Add stirring bar and lid with 3 addition ports (Before you begin step 18.b.) before autoclaving.
	KCl	4.5 g
	MgSO4 anhydrous	0.6 g
	CaCl2 anhydrous	0.076 g
	NaHCO3	1.5 g
	KH2PO4	0.5 g
	FeSO4 7H2O	0.005 g
	Bile salts (no. 3)	0.4 g
	Tryptone	5 g
	Peptone	5 g
	Yeast extract	4.5 g
	Starch (wheat)	5 g
	Pectin (from citrus)	2 g
	Guar gum	1 g
	Xylan (from beech wood)	2 g
	Inulin (from chicory)	1 g
	Arabinogalactan (larch wood)	2 g
	Anti-foam	2.5 mL
	Tween 80	1 mL
2	Porcine gastric mucin (type II)	4 g	200 mL pre-heated 1 M NaOH	Autoclave 121°C, 15 min	Add stirring bar before autoclaving.
3	Casein	3 g	75 mL pre-heated 1.6 M NaOH	Autoclave 110°C, 15 min	Add stirring bar before autoclaving.
4	L-Cysteine HCl	0.8 g	12.5 mL ddH2O	Sterile filtration	–
5	Pantothenate	10 mg	1 mL ddH2O	Sterile filtration	Solve and sterile filter ingredients separately. Combine sterile ingredients in a sterile environment and prepare aliquots (i.e., for 2 L final volume).
	Nicotinamide	5 mg	1 mL ddH2O
	Thiamine	4 mg	1 mL ddH2O
	Biotin	2 mg	1 mL ddH2O, titrated with 1 M NaOH
	Vitamin B12	0.5 mg	1 mL 1 M NaOH
	Menadione	1 mg	1 mL 100% ethanol
	PABA	5 mg	1 mL 95% ethanol
	Hemin	50 mg	2 mL 1.6 M NaOH

Adapted from Macfarlane et al.² and Gibson and Wang.¹²

Table 4.

Exemplary preparation of a cultivation media mimicking infant conditions

Media #	Ingredient	Amount (per L media)	Solved in	Sterilization method	Comments
1	NaCl	4.47 g	300 mL ddH2O	Autoclave 121°C, 15 min	Autoclave in bottle big enough for final volume (i.e., 2 L). Add stirring bar and lid with 3 addition ports (Before you begin step 18.b.) before autoclaving.
	KCl	4.43 g
	MgSO4 anhydrous	1.24 g
	CaCl2 anhydrous	0.1 g
	NaHCO3	1.5 g
	KH2PO4	0.5 g
	FeSO4 7H2O	0.005 g
	Bile salts (no. 3)	0.05 g
	Tryptone	0.5 g
	Peptone	0.5 g
	Yeast extract	2.5 g
	L-Carnitine	0.0015 g
	Tween 80	1 mL
	Anti-foam	2.5 mL
2	Porcine gastric mucin (type II)	4 g	200 mL pre-heated 1 M NaOH	Autoclave 121°C, 15 min	Add stirring bar before autoclaving.
3	WPH	11.2 g	370 mL ddH2O	Autoclave 110°C, 15 min	Dissolve Casein in 1 M NaOH and add it to WPH-formula mix. Adjust pH to 8.0 and add stirring bar before autoclaving.
	Infant formula (Organic Infant Milk PRE, Töpfer GmbH, Germany)	11.88 g
	Casein	0.47 g	25 mL 1 M NaOH
4	L-Cysteine HCl	0.8 g	56 mL ddH2O	Sterile filtration	Solve and sterile filter ingredients together.
	Lactose	5.86 g
	Inositol	0.004 g
5	Pantothenate	10 mg	1 mL ddH2O	Sterile filtration	Solve and sterile filter ingredients separately. Combine sterile ingredients in a sterile environment and prepare aliquots (i.e., for 2 L final volume).
	Nicotinamide	5 mg	1 mL ddH2O
	Thiamine	4 mg	1 mL ddH2O
	Biotin	2 mg	1 mL ddH2O, titrated with 1 M NaOH
	Vitamin B12	0.5 mg	1 mL 1 M NaOH
	Menadione	1 mg	1 mL 100% ethanol
	PABA	5 mg	1 mL 95% ethanol
	Hemin	10 mg	1 mL 1.4 M NH4OH

Adapted from Cinquin et al.³

Selection of fecal sample

Timing: variable

• 11.Use 2–2.5 g fecal material per inoculation for two cultivation vessels (duplicates).

Note: It is possible to use lower amounts but ensure that the same amount is used per inoculation time point.

• 12.Select samples with similar consistency or dry weight and bacterial number to ensure comparability between experiments with different donor samples.

Note: The bacterial number can be determined via anaerobic cultivation on Wilkins-Chalgren Anaerobe (WCA) agar (see step 49).

• 13.Determine the number of cultivation experiments planned with the same donor. Take into account that each cultivation vessel is inoculated twice per experiment.
• 14.Calculate the minimal sample amount and select samples exceeding this amount.
• 15.Ideally, select samples that were quickly processed to reduce O₂ exposure and then stored in 20% glycerol to conserve the bacterial community.

Preparation of solutions, WCA agar, waste bottles and inoculation needles

Timing: 6 h, working time: 4 h

• 16.
  Prepare 1 bottle of 5% H₃PO₄ and 50 mg/mL NaOH per cultivation system.

  • a.
    Consider the pH of cultivation media and target pH during ex vivo cultivation to prepare appropriate volumes of both solutions (i.e., 2 L of 5% H₃PO₄ and 250 mL of 50 mL/mg NaOH if pH needs to be decreased from pH ∼12 to reach a target pH of 6.5).
  • b.
    Prepare lids with 3 addition ports to fit onto reagent bottles.

    • i.
      Prepare blanking ends by tying a knot into ∼10 cm silicone hose pieces (Ø = 2 mm) (see Figure 3).
    • ii.
      On the outside, fit blanking ends on 2 addition ports and one autoclavable air filter (pore size 0.2 μm) connected to 8 cm silicone hose (Ø = 2 mm) on the remaining port.

      Note: Air filters can be secured in the silicone hose with zip ties.

    • iii.
      On the inside, connect ddH₂O rinsed silicone hoses (Ø = 2 mm) with appropriate length to reach the bottle ground to the addition ports covered with blanking ends.
    • iv.
      Fit lids onto respective reagent bottles.
    • v.
      Cover the air filter with aluminum foil.
    • vi.
      Place a clamp on the silicone hoses on the outside of the lid and tighten it.
    • vii.
      Loosely, cover the entire lid with aluminum foil.
  • c.
    Autoclave 121°C, 15 min.
  • d.
    Store at 20°C–25°C for up to 4 weeks.
• 17.
  Prepare 2 L BHI per cultivation system.

  • a.
    Equip the bottles with lids with 3 addition ports (step 17.b.) and autoclave 121°C, 15 min.
  • b.
    Store at 4°C until connecting it to cultivation systems.
• 18.
  Prepare anaerobic WCA agar plates.

  • a.
    Prepare 2.5 ng/mL phenosafranine stock. Sterile filter and store at 20°C–25°C for 2 months.
  • b.
    Prepare L-Cysteine-DTT stock. Sterile filter and store at – 20°C for up to 2 months.
  • c.
    Prepare agar plates and store at 4°C for up to 4 weeks.
  • d.
    Place agar plates into the anaerobic workstation at least 24 h prior to use.
• 19.
  Prepare reduced phosphate-buffered saline (PBS) for inoculation.

  • a.
    Prepare 30 mg/mL L-Cysteine HCl stock. Sterile filter and store at 4°C for up to 4 weeks.
  • b.
    Prepare reduced PBS for inoculation by adding the L-Cysteine HCl stock to sterile PBS. Place it in the anaerobic workstation at least 24 h prior to inoculation and open the lid a half turn.
• 20.Prepare 40% glycerol-PBS stock for aliquoting of ex vivo cultivation samples. Sterile filter and store at 4°C for up to 4 weeks.
• 21.
  Prepare 1 waste bottle with a lid with 4 addition ports per cultivation system.

  • a.
    On the outside, fit blanking ends on 3 addition ports and one autoclavable air filter (pore size 0.2 μm) connected to 8 cm silicone hose (Ø = 2 mm) on the remaining port.

    CRITICAL: Connect the “inlet” side of the filter towards the lid, to decrease the risk for the environment.

  • b.
    On the inside, connect 10 cm silicone hoses (Ø = 3 mm) to the addition ports covered with blanking ends.
  • c.
    Cover air filter with aluminum foil.
  • d.
    Place clamps on the silicone hoses on the outside of the lid and tighten them.
  • e.
    Place lids on empty 5 L bottles, cover the lids with aluminum foil and autoclave 121°C, 20 min.
• 22.Sterilize inoculation needles 24 h before inoculation. Wrap individual inoculation needles in aluminum foil and sterilize at 180°C for 3 h. Prepare 2 needles per cultivation vessel and inoculation time point.

Figure 3.

Picture of blanking ends

Aliquoting of fecal samples in preparation for inoculation

Timing: variable, working time per sample: 1 h

• 23.
  Prepare the sample as follows:

  • a.
    In case of a fresh stool sample:

    • i.
      Determine the weight of the stool sample.
    • ii.
      In a sterile environment, place the container with the stool sample on ice and add 40% glycerol-PBS stock to the sample for a final concentration of 20% glycerol (w/v). Mix well using sterile utensils.
    • iii.
      Aliquot 2.5 g of mixed sample for 2 cultivation vessels or 5 g for 4 cultivation vessels and freeze immediately at – 80°C.
  • b.
    In case of not-aliquoted frozen stool samples:

    • i.
      Clean and disinfect a fume hood before sterilizing with UV light for 30 min.

      Note: UV light can be a health risk. Covering the front of the fume hood with aluminum foil during sterilizing can help.

      Alternatives: It is possible to use a ducted biosafety cabinet instead of a fume hood for this procedure.

    • ii.
      Disinfect the container with liquid N₂, tubes for aliquoting and utensils (from step 24.b.i.) before placing them inside the fume hood. Sterilize with UV light for 15 min.
    • iii.
      Place a clean icebox with dry ice inside the fume hood. Carefully, unwrap the mortar bowl and use the aluminum foil to cover the ice.
    • iv.
      Pour a small amount of liquid N₂ into mortar bowl to cool it down. Use the pestle to spread it around to create an anaerobic environment.
    • v.
      Remove the fecal sample from dry ice and disinfect the outside before introducing it into the fume hood. Add the entire sample to the mortar bowl.
    • vi.
      Regularly, add liquid N₂ to keep sample frozen and anaerobic.

      CRITICAL: Sample consistency should be frozen or smooth but never liquid.

    • vii.
      Use the pestle to homogenize the sample with round motions.
    • viii.
      Aliquot the desired amount into tubes and store on dry ice immediately.
    • ix.
      Repeat steps 24.b.i. – 24.b.ix. to aliquot another fecal sample. Use a new set of clean instruments for each sample to avoid cross-contamination.
    • x.
      Disinfect all used utensils and clean them. Sterilize the fume hood with UV light for 90 min.

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Biological samples
Infant feces	Infantibio-II Study	N/A
Adult feces	BISS Study	N/A
Chemicals, peptides, and recombinant proteins
Sodium chloride	Sigma	Cat#31434-M
Potassium chloride	Sigma	Cat#P9333
Magnesium sulfate anhydrous	Sigma	Cat#746452
Calcium chloride anhydrous	Sigma	Cat#C1016
Sodium hydrogen carbonate	Sigma	Cat#S6014
Potassium dihydrogen phosphate	Sigma	Cat#P5655
Iron-(II)-sulfate heptahydrate	Fisher	Cat#201392500
Bile salts (no. 3)	Fluka	Cat#48305
Tryptone	Sigma	Cat#T9410
Peptone	Sigma	Cat#91249
Yeast extract	Sigma	Cat#Y1625
Starch (from wheat)	Sigma	Cat#S5127
Pectin (from citrus)	Alfa Aesar	Cat#J61021
Guar gum	Sigma	Cat#G4129
Xylan (from beech wood)	Roth	Cat#4414.4
Inulin (from chicory)	Sigma	Cat#I2255
Arabinogalactan (from larch wood)	SCBT	Cat#sc-210833
Anti-foam solution B	Sigma	Cat#A5757
Tween 80	Sigma	Cat#P8074
Porcine gastric mucin (type II)	Sigma	Cat#M2378
Casein	Fisher	Cat#A13707
Hydrolyzed whey protein, pre-digested, unflavored	MYPROTEIN	N/A
Organic infant milk PRE	Töpfer GmbH, Germany	N/A
L-cysteine HCl	Fisher Acros	Cat#111781000
Lactose	Fisher	Cat#36889
Inositol	Sigma	Cat#57570
L-carnitine	Sigma	Cat#8.40092
Pantothenate	AppliChem	Cat#A2088
Nicotinamide	VWR	Cat#A15970
Thiamine	AppliChem	Cat#A2088
Biotin	Fisher	Cat#23009
Vitamin B12	Sigma	Cat#V2876
Menadione	Fisher	Cat#127180250
4-Aminobenzoic acid (PABA)	Sigma	Cat#A9878
Hemin	Sigma	Cat#51280
Phosphoric acid, 85.5%	Fisher	Cat#11478443
Sodium hydroxide pellets	Sigma	Cat#S5881
Brain-heart infusion	Sigma	Cat#53286
WCA broth	Th. Geyer	Cat#CM0643B
Agar agar	Fisher Scientific	Cat#10572775
Phenosafranine	Sigma	Cat#199648
L-cysteine	Roth	Cat#3467.x
DL-dithiothreitol (DTT)	Sigma	Cat#43819
Phosphate-buffered saline	Invitrogen	Cat#18912-014
Glycerol, 100%	Fisher	Cat#BP229-4
Buffer solution, pH 4.01	VWR	Cat#wtwa 109110
Buffer solution, pH 6.87	VWR	Cat#662-0326
Buffer solution, pH 9.18	VWR	Cat#662-0327
Buffer solution, redox 271 mV	VWR	Cat#HAMI238228
Buffer solution, redox 475 mV	VWR	Cat#HAMI238227
Sodium hydroxide solution, 1 M , 10 L	Fisher	Cat#J762024
Ethanol, absolute	VWR	Cat#20821.321
NH4OH, ≥ 25%	Sigma	Cat#30501-M
KCl, 3 M, 1 L	VWR	Cat#83605.290
Software and algorithms
Eve® Bioprocess Platform Software	Infors HT	Cat#512616
IMNGS2	N/A	https://imngs2.org
Rhea	Langkouvardos et al.13	https://github.com/Lagkouvardos/Rhea
RStudio	RStudio	https://posit.co/products/open-source/rstudio/; RRID: SCR_000432
Cosinor-fitting	Cornelissen et al.14	N/A
Other
Multifors 2 Microbial 1.4 L TV with top plate NW90, 4 cultivation vessels, gas strategy N2	Infors HT	Cat#503078
Hose nipple for reagent bottle inlet	Infors HT	Cat#28885
Axial marine-type impel D = 50 × 5 left	Infors HT	Cat#62027
Harvest pipe D = 6 mm NW90 flat bottom	Infors HT	Cat#62376
Clamping device 05 mm Exzentrr. PG13.5	Infors HT	Cat#67613
Autoclavable pump head ID = 2.5 mm	Infors HT	Cat#28722
Reagent bottle 250 mL 3 + 1 connections	Infors HT	Cat#29021
Super Safe Sampler	Infors HT	Cat#80836
pH-sensor EasyFerm Plus Arc 225 mm	Hamilton	Cat#238633-1343
Redox-sensor EasyFerm Plus ORP Arc 225	Hamilton	Cat#243051
Vacuum grease	Sigma-Aldrich	Cat#18405
Magnetic stirring bars, double ended	VWR	Cat#442-0478
Low-temperature thermostat (i.e., Ecoline RE104)	Lauda	N/A
Silicone hose 2 × 6 mm, 25 m	VWR	Cat#228-1451
Silicone hose 3 × 6 mm, 25 m	VWR	Cat#228-1517
Distributors for bottles	VWR	Cat#D606-08
Midisart 2000, pore 0.2 μm, sterile	Th. Geyer	Cat#17805----G
Minisart RC25 syringe filter, pore 0.2 μm	Th. Geyer	Cat#17764----K
Syringe, Luer Lock, 5 mL	Fisher	Cat#14955458
Syringe, Luer Lock, 10 mL	Fisher	Cat#14955459
Syringe, Luer Lock, 60 mL	Fisher	Cat#15899152
Arium Mini Essential Lab Water Purification System	Sartorius	Cat#H20-MU-UV-T
Cell strainer, sterile, mesh size 70 μm	Fisher	Cat# 11597522
Cannula for inoculation (Luer-Lock connection, 2.0 × 200 mm)	neoLab	Cat#2-3113
Whitley A35 anaerobic workstation (90% N2, 10% H2)	MeintrupDWS	N/A
Ceramic mortar, 75 mL	VWR	Cat#410-0109
Ceramic pestel, 30 mm	VWR	Cat#140-0120
96-well plate (U-bottom, sterile, untreated)	Sarstedt	Cat#82.1582.001
DNA-Stool stabilizer	Invitek	Cat#1038111100
RNAlater	Sigma	Cat#R0901

Materials and equipment

5% H₃PO₄

Reagent	Final concentration	Amount
85.5% Phosphoric acid	5%	58.5 mL
ddH2O	N/A	941.5 mL
Total	N/A	1000 mL

Autoclave and store at 20°C–25°C for up to 4 weeks.

50 mg/mL NaOH

Reagent	Final concentration	Amount
NaOH pellets	50 mg/mL	50 g
ddH2O	N/A	1000 mL
Total	N/A	1000 mL

Autoclave and store at 20°C–25°C for up to 4 weeks.

L-Cysteine-DTT stock

Reagent	Final concentration	Amount
L-Cysteine	50 mg/mL	2.5 g
DTT	20 mg/mL	1 g
ddH2O	N/A	50 mL
Total	N/A	50 mL

Sterile filter and store at – 20°C for up to 2 months.

2.5 ng/mL phenosafranine stock

Reagent	Final concentration	Amount
Phenosafranine	2.5 ng/mL	62.5 mg
ddH2O	N/A	50 mL
Total	N/A	50 mL

Sterile filter and store at RT for 2 months.

Anaerobic Wilkins-Chalgren Anaerobe (WCA) agar

Reagent	Final concentration	Amount
WCA broth	N/A	33 g
Agar	N/A	22.5 g
ddH2O	N/A	1000 mL
2.5 ng/mL phenosafranine stock	2.5 ng/mL	2 mL
L-Cysteine-DTT stock	0.5 mg/mL L-Cysteine, 0.2 mg/mL DTT	10 mL
Total	N/A	1012 mL

Store agar plates at 4°C for up to 4 weeks.

Note: Mix WCA broth, agar and ddH₂O, adjust pH to 7.1, add phenosafranine stock and autoclave, 121°C for 15 min. Cool down to 50°C in a water bath, add L-Cysteine-DTT stock in sterile environment and mix well. Pour 20 mL in sterile petri dishes and let dry.

30 mg/mL L-Cysteine HCl stock

Reagent	Final concentration	Amount
L-Cysteine HCl	30 mg/mL	150 mg
ddH2O	N/A	5 mL
Total	N/A	5 mL

Sterile filter and store at 4°C for up to 4 weeks.

Reduced PBS for inoculation

Reagent	Final concentration	Amount
30 mg/mL L-Cysteine HCl stock	0.3 mg/mL	500 μL
PBS, sterile	N/A	50 mL
Total	N/A	50.5 mL

Store at 4°C for up to 4 weeks.

40% Glycerol-PBS Stock

Reagent	Final concentration	Amount
100% glycerol	40%	400 mL
PBS	N/A	600 mL
Total	N/A	1000 mL

Sterile filter and store at 4°C for up to 4 weeks.

Note: To expedite the reduction, 0.2 mg/mL peptone and 0.5 mg/mL L-Cysteine can be added.

Material for aliquoting of fecal samples in preparation for inoculation.

• •
  For fresh samples:

  • ○
    Spoons, individually wrapped in tinfoil, autoclaved (121°C, 20 min).
  • ○
    Sterile 40% glycerol-PBS stock with 0.2 mg/mL peptone and 0.5 mg/mL L-Cysteine.
• •For frozen samples: ceramic mortars and pestles, spatulas and spoons, individually wrapped in tinfoil, autoclaved (121°C, 20 min).

Material for preparation of one inoculum (1 donor sample including spare). Place material into anaerobic workstation at least 24 h before preparation of the inoculum.

• •Reduced PBS for inoculation.
• •4× WCA agar plates.
• •3 × 10 mL syringes.
• •3 × 24 G cannulas, preferably blunt.
• •2 × 50 mL Falcon tubes.
• •2 × 70 μm cell strainers.
• •3× autoclaved 1.5 mL microtubes.
• •1 box each of autoclaved 1000 μL, 100 μL and 10 μL tips.
• •1× 96-well plate (U-bottom, untreated).

Material for collecting samples from ex vivo cultivation systems (1 system).

• •1 Luer Lock syringe, 5 mL, unlabeled.
• •1 Luer Lock syringe, volume depending on desired sampling volume, labeled with vessel number immediately before sample collection.
• •Sterile, labeled microtubes or Falcon tubes, depending on desired sampling volume.
• •If applicable, sterile 40% Glycerol-PBS Stock, DNA stabilizer or RNA later.

Step-by-step method details

Setup of ex vivo cultivation systems

Timing: 4–5 days (depending on number of cultivation vessels)

General setup of ex vivo cultivation systems

Timing: 2 h for 1 cultivation system (2 vessels)

The cultivation vessels are set up according to the manufacturer’s instructions (Multifors operating user’s guide for microorganisms).

CRITICAL: Apply vacuum grease to all O-rings during setup to ensure they seal tightly. This prevents loss of pressure and minimizes the risk of contamination throughout the experiment.

• 1.Mount bearing holder, addition port adapters, hose holders, stirrer shafts, baffles, impellers, flow deflectors, magnetic couplings and immersion pocket according to the manufacturer’s instructions.

Note: These parts can stay mounted after the end of an experiment.

• 2.
  Mount the exit gas cooler onto the vessel top plate according to the manufacturer’s instructions with the following modifications:

  • a.
    Equip the exit gas filter (pore size 0.2 μm) with 15 cm tube (Ø = 3 mm) on the outlet side of the filter.
  • b.
    Plug the inlet side of the filter into the piece of hose on the exit gas cooler and secure it.
  • c.
    Do not cover the filter with aluminum foil before autoclaving.
• 3.
  Mount the sparger according to the manufacturer’s instructions. Prepare the connection to the basic unit as follows:

  • a.
    Equip a sterile filter (pore size 0.2 μm) with 2 pieces of hose (Ø = 3 mm; 15 cm). Push the hose onto the filter until the 2^nd prong on both sides.
  • b.
    Remove the plug from the N₂ supply in the basic unit and fit it into the hose on the inlet side of the filter.

    • i.
      Cover the plug with a small piece of aluminum foil.
    • ii.
      Add one clamp to the hose close to the plug and tighten it.
  • c.
    Fit the hose on the outlet side of the filter to the sparger and secure it with a cable tie. Add 2 clamps to the hose close to the sparger and tighten them.
• 4.
  Mount the dip tube and prepare the Super Safe Sampler according to the manufacturer’s instructions. Mind the following points:

  • a.
    Adjust the height of the dip tube to half of the working volume (i.e., 200 mL for a working volume of 400 mL).
  • b.
    Use ddH₂O rinsed silicone hose (Ø = 2 mm) to connect dip tube and sampler.
  • c.
    Only use filters (pore size 0.2 μm) suitable for autoclaving.
  • d.
    Add a clamp to the hose close to the dip tube and tighten it.

CRITICAL: Check that you are able to collect samples with the Super Safe Sampler before connecting it to the dip tube (i.e., by sampling ddH₂O).

• 5.
  Mount the waste pipe in the same way as the dip tube.

  Note: Before mounting, ensure that waste and dip tube are similar to set up.

  • a.
    Adjust the height of the waste pipe to the working volume (i.e., 400 mL).
  • b.
    Fit ddH₂O rinsed silicone hose (Ø = 3 mm) onto waste pipe and secure it with a cable tie.

    Note: The length of the hose depends on the location of the waste bottle.

  • c.
    Cover the end of the hose in aluminum foil.
  • d.
    Place a clamp on the hose close to the waste pipe and tighten it.
  • e.
    For easy transport, roll up the hose and cover it in aluminum foil. Make sure that the end of the hose is still covered.

    Note: If the slotted screw on the clamping adapter is inaccessible after screwing the clamping adapter into the vessel top plate, adding a septum collar in between top plate and clamping adapter can help.

• 6.Equip one port on the vessel top plate with a septum and septum collar for inoculation according to the manufacturer’s instructions. Mount a blanking plug (Ø = 12 mm) into the septum collar.

Note: For easy access during the inoculation, use a port at the front of the vessel top plate.

• 7.Fasten the clamping ring around the vessel top plate according to the manufacturer’s instructions.
• 8.
  Prepare pumps and hoses for the addition of acid, base and feed.

  • a.
    Fit 3 pumps per vessel (acid, base and feed) on the mounting plate on the basic unit.
  • b.
    Prepare 3 pieces of ddH₂O rinsed silicone hose (Ø = 2 mm) to connect the pumps to the addition ports on the vessel top plate.

    • i.
      Choose the length of the silicone hoses to fit between pumps and addition ports without any tension.
    • ii.
      Connect the silicone hoses to the hose connectors on the right side of the pumps.

      Note: The pumps rotate counter-clockwise.

    • iii.
      Secure both sides of the hose connector with cable ties.
    • iv.
      Connect the silicone hoses to addition ports on the vessel top plate and secure them with cable ties.

      Note: To facilitate monitoring of the media influx during the experiment, connect the silicone hose from the feed pump to the addition port in the front.

  • c.
    Prepare 3 pieces of ddH₂O rinsed silicone hose (Ø = 2 mm) to connect the pumps to the reagent bottles.

    • i.
      Chose the length of the silicone hoses to fit between pumps and reagent bottles without any tension.

      Note: Ensure that the silicone hose connected to the media bottle is long enough to move around when connecting a new media bottle.

    • ii.
      Connect the silicone hoses to the hose connectors on the left side of the pumps.
    • iii.
      Secure both sides of the hose connector with cable ties.
    • iv.
      Cover the ends of the silicone hoses (that will connect to the reagent bottles) with aluminum foil.
    • v.
      Place a clamp on the hose close to the aluminum foil.
    • vi.
      Roll up each hose and cover them with aluminum foil.
  • d.
    Prepare 1 piece of silicone hose (Ø = 2 mm) of approx. 5 cm.

    • i.
      Fit it on the remaining addition port on the vessel top plate.
    • ii.
      Secure it with a cable tie and place a clamp on the end of the hose to close this port.
• 9.Mount blanking plugs (Ø = 12 mm) into any port that is not used according to the manufacturer’s instructions.

CRITICAL: Ensure that the stirrer shaft tip is inserted into the centering bearing (troubleshooting 1).

Note: Arrange the appliances in the vessel top plate in such a way that it matches the setup of the basic unit of the cultivation system (i.e., mount the sparger close to the N₂ outlet, the exit gas cooler close to the rapid couplings or the electrode holders close to the cable connections for the electrodes) (see Figure 4).

Note: It is possible to mount all accessories on the top plate during one session and continue with adding the silicone hoses at a later time point. In case of a longer pause (i.e., longer than overnight), cover all openings with aluminum foil to avoid dust and small particles from entering the cultivation vessels.

Pause Point: The setup can be paused at any time for as long as necessary if all openings are covered (i.e., using blanking plugs or aluminum foil.

Figure 4.

Description of the advised setup for vessel top plates

(A) Picture of vessel top plates of one cultivation system with the advised setup.

(B) Schematic depiction of the advised setup on vessel top plates.

Preparing ex vivo cultivation systems for autoclaving and autoclaving

Timing: 3 h for 1 cultivation system, working time: 1 h

Note: Only continue with these steps immediately before autoclaving.

CRITICAL: Use ddH₂O for all solutions inside the cultivation vessels, including during autoclaving, media preparation and cleaning.

• 10.Remove the blanking plugs (Ø = 12 mm) from the vessel top plate.
• 11.Use one of the empty ports to add ddH₂O to the vessel until the working volume is reached.
• 12.
  Mount the sensor holders for pH and redox sensor on the vessel top plate.

  • a.
    Insert greased O-rings into each port and insert hollow screws on top.
  • b.
    Install the fork of the guide bar at the bottom of the hollow screw and screw it down tightly using a hex socket wrench.
  • c.
    Fit the support guide onto the guide bar.
  • d.
    Ensure that the holes of hollow screw and support guide overlap and tighten the grub screw of the support guide with the key.
• 13.
  Calibrate one pH and one redox sensor per vessel using the appropriate buffers (troubleshooting 2).

  • a.
    Insert the sensors into the sensor holders after successful calibration and screw them down tightly.
  • b.
    Repeat with the second vessel of the same system.
• 14.Remove the mounting plate with pump heads from the drive shafts on the basic unit and fit it in the vessel holder.
• 15.Tighten the bracket between the thermal blocks on the basic unit to release the vessel holder.
• 16.Make sure that the vessels are not connected to the basic unit in any way before lifting the vessel holder up to remove it from the basic unit.
• 17.
  Place the vessel holder into the autoclave. Ensure that:

  • a.
    The electronic ends of sensors are not in contact with aluminum foil or metal clamps.
  • b.
    The silicone hoses on the filters of the exit gas coolers are not bended strongly or otherwise obstructed.
  • c.
    The silicone hoses connected to the spargers are not bended strongly to minimize the risk of rupture during autoclaving and the end points up to reduce inflow of water into the silicone hose.
  • d.
    The vessel holder does not stand on any silicone hoses.
• 18.Autoclave the cultivation system: 121°C, 15 min

Setup after autoclaving, preparation for sterility test

Timing: 6 h for 1 cultivation system, working time: 1 h

• 19.Immediately after opening the autoclave, place clamps on the silicone hoses on the filters of the exit gas coolers while still in the autoclave and tighten the clamps to close the systems.

CRITICAL: Check that all silicone tubes are intact, the filter is still in the exit gas cooler and that there is no water in the silicone hose on the inlet side of the filter of the gas entry (troubleshooting 3). Check the magnetic coupling for rust (troubleshooting 4).

• 20.Place the vessel holder into the basic unit from above and release the bracket between the thermal blocks to secure the vessels.
• 21.Mount the mounting plate with the pump heads on the drift shafts on the basic unit.
• 22.Insert temperature sensor into the immersion pocket on the top plate.
• 23.Connect the pH and redox sensors to the basic unit. Check the output on the touch screen.
• 24.Connect the exit gas cooler to the basic unit according to the manufacturer’s instructions and adjust the water flow to the highest flow.
• 25.
  Connect the waste tube to the waste bottle as follows:

  Note: Connect both vessels of one cultivation system to the same waste bottle.

  • a.
    Carefully remove the aluminum foil covering the rolled up silicone hose.

    CRITICAL: Ensure that the end of the hose remains covered in aluminum foil during this process.

  • b.
    Remove the aluminum foil covering the lid of the waste bottle (troubleshooting 5).
  • c.
    Disinfect the lid, the end of the silicone hose, blanking ends and gloves to minimize the risk of contamination.
  • d.
    Prepare the waste bottle for connection by pushing the blanking end upwards until only a small part of the addition port is still covered and it can be easily removed.
  • e.
    Remove the blanking end from the addition port, take off the aluminum foil covering the end of the silicone hose and immediately push the hose onto the addition port.
  • f.
    Remove the clamp from the silicone hose.

    CRITICAL: Due to slight under pressure in the system after autoclaving, air may be drawn into the vessel through the waste tube. Ensure that waste bottles are sterile before connecting them to waste tubes.

    Optional: Waste bottles can be placed into a fume hood over the course of the experiment to minimize unpleasant smell.

• 26.
  Connect the sparger to the basic unit as follows:

  • a.
    Set the N₂ supply to the basic unit to 2.5 bar.
  • b.
    Use the touch screen to set the N₂ influx into the vessel to 2 L/min.
  • c.
    Remove the aluminum foil from the plug in the silicone hose on the inlet side of the filter and push the plug into the N₂ supply of the basic unit.
  • d.
    Open the clamp between the plug and filter.
  • e.
    Check if water rose into the silicone hose between sparger and filter. If so, proceed with care to prevent water from reaching the filter and blocking it.
  • f.
    Open and remove the lowest clamp between sparger and filter, carefully, and monitor the water level in the silicone hose.

    • i.
      The water level is stable: Carefully remove the second clamp. Water will be pushed through the waste tube to release pressure.
    • ii.
      The water level is rising towards the filter: Place a clamp over the remaining clamp towards the filter and close it loosely. Open the lower clamp very carefully and monitor the water level. If it rises, close the upper clamp completely and start this step (26.f.) over.
  • g.
    Use the touch screen to set the N₂ influx into the vessel to 0.01 L/min (troubleshooting 6).
• 27.
  Use the touch screen to adjust the following settings:

  • a.
    Start temperature control; set point: 37°C (troubleshooting 7).
  • b.
    Start stirrer; set point: 40 rpm (troubleshooting 8).
  • c.
    Stop pH control if it was turned on.
• 28.
  Connect the acid, base and media bottles to the respective pumps.

  • a.
    Follow the same procedure as for connecting the waste tube to the waste bottle (step 25).
  • b.
    Ensure that the pumps are connected to the correct bottles.
  • c.
    Place the media bottle on a magnetic stirrer and start stirring.
  • d.
    Fill the silicone hoses by pressing the rocker switch to the right side.

    CRITICAL: Pressing the rocker switch to the left side moves the solution from the addition port into the reagent bottles. This may cause contaminations in the reagent bottles.

  • e.
    Consecutively, add 5 drops of acid and base by pressing the rocker switch to the right and monitor the pH on the touch screen. Only continue when the pH changes upon addition (troubleshooting 9).
• 29.
  Exchange the ddH₂O in the vessel for BHI:

  Note: To ensure complete exchange, add the working volume twice per vessel (i.e., 800 mL with a working volume of 400 mL).

  • a.
    Use the touch screen to set the feed rate to an appropriate value (i.e., 100% for the fastest exchange).

    Note: It is possible to exchange for BHI slowly (i.e., overnight). In this case, ensure that the feed rate is appropriate to prevent the bottle from running empty.

  • b.
    Use the touch screen to stop the feed after the necessary volume was added to the vessels.

Sterility test of the cultivation vessels

Timing: 48 h independent of number of cultivation systems, working time: 10 min

• 30.
  Check that only the following settings are turned on during the sterility test:

  • a.
    N₂ influx (set point: 0.01 L/min)
  • b.
    Temperature (set point: 37°C)
  • c.
    Stirrer (set point: 40 rpm)
• 31.
  Incubate the vessels for 48 h.

  • a.
    BHI looks clear: The vessel is sterile and the experiment can start (continue with step 32).
  • b.
    BHI looks cloudy: The vessel is contaminated.

    • i.
      Remove the BHI from the vessel, rinse the vessel and pipes with ddH₂O.
    • ii.
      Add ddH₂O until the working volume is reached.
    • iii.
      Prepare both vessels of the affected system for autoclaving (continue from step 14).

      CRITICAL: Remove the clamp from the silicone hose on the filter of the exit gas cooler before autoclaving. This is the only pressure relief for the vessel during autoclaving.

Note: The length of the incubation with BHI can be prolonged if necessary (i.e., due to a contamination of a single vessel or when autoclaving systems over multiple days) but it should not be shortened. Consider stopping N₂ influx if the incubation is prolonged to decrease experimental costs.

Pause Point: The incubation with BHI can be extended to 7 days.

Preparing the cultivation vessels for inoculation

Timing: 36 h independent of number of cultivation systems, working time: 30 min per system

• 32.
  Exchange BHI for the cultivation media.

  Note: To ensure complete exchange, add the working volume twice per vessel (i.e., 800 mL with a working volume of 400 mL).

  • a.
    Connect the feed pump to the bottle containing cultivation media:

    • i.
      Place a clamp on the silicone hose connected to the bottle with BHI.
    • ii.
      Disinfect the lids of both media bottles (BHI and cultivation media), the silicone hose, blanking ends and gloves.
    • iii.
      Push the silicone hose upwards until only a small part of the addition port is covered and it can be easily removed. Repeat with the blanking end on the bottle with cultivation media.
    • iv.
      Remove the blanking end and the end of the hose from the addition ports and immediately push the hose onto the bottle with cultivation media.
    • v.
      Remove the clamp.
    • vi.
      Repeat this for the second vessel.
  • b.
    Set the feed rate to an appropriate value (see step 29.a).
  • c.
    Use the touch screen to start the pH control (for the choice of an appropriate set point see before you begin step 2).

    Note: Exchange the media approximately 36 h before the inoculation to allow for sterility testing. Do not exchange it earlier than 48 h to ensure freshness of media and not later than 24 h before inoculation for sterility test.

• 33.Change the phase in eve® software (see Manufacturer’s instructions for details).
• 34.
  After media exchange, check sterility of cultivation media in the cultivation vessels.

  • a.
    Carefully, remove the aluminum foil from the Super Safe Sampler.
  • b.
    Collect a sample from each cultivation vessel (see steps 38–42).
  • c.
    In a sterile environment, plate undiluted cultivation media on WCA agar and incubate it anaerobically (37°C, 90% N₂, 10% H₂) for min. 24 h.

Preparation of cultivation media

Timing: 8 h, working time: 4 h

The cultivation media should be prepared 24 h before it is needed in parallel to the above-mentioned steps. Store media at 4°C until needed and warm up to RT before connecting to the cultivation system to minimize the risk of blockages with high viscosity media.

• 35.Prepare one bottle of cultivation media for one cultivation system.
• 36.Depending on the retention time, prepare cultivation media in volumes allowing exchange of media bottles during working hours (i.e., 1.8 L for 12.5 h retention time with 400 mL working volume allowing a change of media bottles every 24 h).
• 37.Prepare the cultivation media as tested during the experimental media design.

Collecting samples from ex vivo cultivation systems

Timing: 30 min per cultivation system (+ 15 min for each additional cultivation system)

Samples can be collected from the cultivation vessels at any time. Using super safe samplers can reduce the risk of contamination.

• 38.
  Preparations before collecting a sample:

  • a.
    Place tubes for aliquoting in a sterile environment.
  • b.
    Prepare dry ice for snap freezing of samples after aliquoting.
  • c.
    Prepare 2 Luer Lock syringes per vessel. Label one syringe per vessel to distinguish during aliquoting.
  • d.
    Remove syringes from the hose on the filter of the Super Safe Sampler, fill with air and re-insert them into the hose.
  • e.
    Disinfect the sample valves.
• 39.
  Collect sample from one cultivation vessel as follows:

  • a.
    To discard liquid leftover in the sampling tube and disinfectant residues on the sample valve, screw the unlabeled Luer Lock syringe on the sample valve, open the clamp on the hose and draw up ∼3 mL of cultivation media (troubleshooting 10).
  • b.
    Remove the syringe from the sample valve, discard it and immediately connect the labeled Luer Lock syringe.
  • c.
    Draw up the desired sample volume.
  • d.
    Rinse the dip tube with sterile air by pushing the piston of the syringe filled with air.
  • e.
    Remove residual fluid from the Super Safe Sampler by pushing in sterile air and simultaneously pulling the piston of the Luer Lock syringe (troubleshooting 11).
  • f.
    Close the clamp on the hose to the dip tube.
• 40.Repeat step 39 for the remaining cultivation vessels, consecutively.

CRITICAL: Only remove the Luer Lock syringe from the sample valves after collecting samples from all desired cultivation vessels to avoid prolonged exposure to air.

• 41.Remove the Luer Lock syringes from the sample valves and aliquot the liquid in a sterile environment. Snap freeze aliquots immediately according to experimental design.

Note: Consider storing samples for gDNA isolation in DNA Stabilizer (INVITEK) for long-term storage.

• 42.Disinfect the sample valves.

Preparation of fecal samples and inoculation into ex vivo cultivation systems

Timing: 2.5 h for 1 donor sample, working time: 1.5 h

To inoculate the cultivation vessels with the donor sample, solid particles need to be removed from the sample by straining and diluting in reduced PBS.

CRITICAL: Before inoculation, consider labeling the cultivation vessels according to the tested conditions (i.e., different cultivation media, donor samples) to avoid mix-ups later on.

• 43.
  Check before preparing the inoculum:

  • a.
    The N₂ influx is sufficient.
  • b.
    The volume and pH of media in the cultivation vessels are correct (troubleshooting 12, 13, and 14).
  • c.
    The plates with cultivation media (see step 34.c) show no sign of contamination.
  • d.
    There are 2 sterilized inoculation needles for each cultivation vessel available (see step 50).
• 44.Thaw fecal sample(s) on ice.

CRITICAL: Plan the thawing time according to the sample amount.

• 45.
  Prepare the vessels for inoculation:

  Note: This can be done by a second person while the inoculum is prepared.

  • a.
    Use the touch screen to stop pH control and feed. For other parameters, apply the set points listed in step 30.
  • b.
    Increase the height of the waste pipe by 50 mL to avoid any loss of cultivation media after inoculation.

    CRITICAL: Only perform this step after the feed was stopped.

  • c.
    Remove the blanking plugs from the septum collars while holding on to the septum collar.
  • d.
    Apply disinfectant onto the septum membranes.
• 46.
  Prepare the inoculum as follows:

  Note: All steps are carried out anaerobically (i.e., in an anaerobic workstation).

  • a.
    Place 70 μm sterile cell strainer in 50 mL sterile Falcon and add thawed sample in the strainer.
  • b.
    Use a syringe piston to sift sample through the strainer, add reduced PBS for inoculation as needed.
  • c.
    Prepare aliquots for gDNA isolation (i.e., 600 μL) and plating (i.e., 30 μL).
  • d.
    Divide the remaining volume into syringes using cannulas.

    Note: Prepare one syringe per vessel that will be inoculated.

  • e.
    Add anaerobic atmosphere in the syringe to ensure no liquid remains in the cannula.
  • f.
    Remove the syringes with attached cannulas from the anaerobic workstation and immediately proceed with the inoculation.
• 47.
  Inoculate the vessels as follows:

  • a.
    Remove the cannula from the first syringe and immediately replace it with the needle for inoculation.
  • b.
    Pierce the septum membrane of the first cultivation vessel in the middle, insert the needle into the medium and pour in the sample (troubleshooting 15).

    CRITICAL: Hold on to the part connecting the inoculation needle and the syringe containing the inoculum to avoid pressing the syringe out of the needle and spilling the inoculum. This can be a health risk. Avoid touching the stirrer with the needle to prevent bending the needles and scratching the stirrer.

  • c.
    Draw up media into the syringe and pour it back. Repeat this step twice.
  • d.
    Draw up atmosphere into the syringe and pour it back to remove droplets in needle.
  • e.
    Remove needle.
  • f.
    Repeat steps b)-f) to inoculate the remaining vessels.
  • g.
    Click “Inoculate” and insert the inoculated volume [mL] per vessel in eve® software.
  • h.
    Apply disinfectant onto septum membranes and screw down blanking plugs into septum collars.
  • i.
    Freeze the aliquot for gDNA isolation at – 80°C.
• 48.In case multiple donor samples shall be used for inoculation, repeat steps 45–47 consecutively, to avoid cross-contamination between the samples.

Note: The second donor sample can thaw during preparation of the first inoculum.

• 49.
  After all bioreactors are inoculated with the respective donor sample, prepare 10⁻¹ to 10⁻⁸ dilutions of the inoculum in reduced PBS

  • a.
    Plate them in triplicates on WCA agar in the anaerobic workstation.
  • b.
    Incubate for 72 h and count the colony forming units (CFU) daily.
  • c.
    Calculate the number of cultivable bacteria (CFU/mL) per dilution.

    $$Cultivablebacteria\left(\frac{CFU}{mL}\right)=\frac{CountedCFU\times Dilution}{platedvolume\left(mL\right)}$$
  • d.
    Determine the mean bacterial number per inoculum.
• 50.
  Clean the inoculation needles with ddH₂O, 70% ethanol and supplied wires for cleaning.

  Note: Liquid waste from cleaning the needles needs to be autoclaved before disposal.

  • a.
    Dry the needles using clean pressurized air.
  • b.
    Wrap individual needles in aluminum foil and sterilize them at 180°C for 3 h.

Batch cultivation of fecal material

Timing: 48 h independent of number of cultivation systems, working time: 2 h for 1 donor sample

The batch cultivation allows the bacterial community to colonize the cultivation vessels before the start of the continuous cultivation. A second inoculation 24 h after the first one improves the transfer of zOTUs from the donor sample into the cultivation vessel (see Figure 5).

• 51.
  24 h after the first inoculation, increase the media volume in the cultivation vessels to double the working volume (i.e., 800 mL with a working volume of 400 mL).

  • a.
    Connect a bottle of fresh media (see step 32.a.).
  • b.
    Adjust the height of the waste pipe to double the working volume + 50 mL to avoid loss of cultivation media.
  • c.
    Use the touch screen to start pH control and feed at 100% feed rate.
  • d.
    Stop the feed using the touch screen when the cultivation media reaches double the working volume.
  • e.
    Stop the pH control using the touch screen when the set point is reached.

Note: If necessary, further increase the height of the waste pipe to avoid loss of cultivation media.

• 52.Inoculate for the second time as soon as the pH reaches the set point. Refer to steps 44–50 for inoculum preparation and inoculation.

CRITICAL: Time the preparation of the inoculum to finish when the pH set point is reached.

Figure 5.

Influence of the inoculation strategy on transfer of zOTUs from adult donor sample into cultivation vessels

(A) Two inoculations at t = 0 h and t = 24 h of batch culture increased the richness and transfer efficiency after 7 days of continuous culture.

(B) Addition of an SCFA mix (1000 μM acetate, 333 μM propionate, 333 μM butyrate) did not improve richness and transfer efficiency. Richness represented as mean of two technical replicates ± SD. Transfer efficiency represented as mean of two technical replicates. n = 2.

Continuous cultivation

Timing: up to 19 days, depends on research question

During the continuous cultivation, media is constantly fed into the cultivation system and excess volume removed to mimic intestinal conditions. It is important to monitor the conditions (i.e., media volume, N₂ influx, pH, redox potential) closely throughout this phase.

• 53.
  48 h after the first inoculation, decrease the media volume in the cultivation vessels to the working volume (400 mL).

  • a.
    Collect samples from each cultivation vessel (see steps 38–42).

    Note: In this step, it is possible to collect large sample volumes before the media volume is reduced to the working volume.

  • b.
    Spray waste tube with disinfectant in excess. Slowly, adjust the height of the waste pipe to the working volume (troubleshooting 13).

    CRITICAL: Turbidity of the cultivation media might impair the visibility of the waste pipe. To not remove more cultivation media than needed, proceed slowly and with care.

• 54.Use the touch screen to start the pH control and feed rate with an appropriate set point (for the choice of an appropriate set point see before you begin step 2).
• 55.Change the phase in eve® software.
• 56.Ensure constant media, acid and base supply by exchanging media bottles before they are empty (see step 32.a.).

CRITICAL: Regularly check the media supply by looking into the vessels and observing media dropping in from the addition port in the vessel top plate (troubleshooting 12).

• 57.Regularly check that the N₂ pressure is sufficient (troubleshooting 6).
• 58.Collect samples according to the experimental design (see before you begin step 2.d.).
• 59.Regularly check for any abnormalities in eve® software. Pay attention to alarms given by the software and on the touch screen of the basic unit.
• 60.For rhythmicity sampling: Collect samples at least every 4 h for 48 h after the bacterial communities stabilized on day 7 of media exposure (see before you begin step 2.d).
• 61.
  For continuous cultivation with different cultivation media:

  • a.
    If applicable: Perform rhythmicity sampling before the change of media (see step 60).
  • b.
    On the last day of media 1, collect a sample from each vessel (see steps 38–42).
  • c.
    Connect bottles containing media 2 to all vessels (see step 32.a).
  • d.
    Change the phase in eve® software.

Note: The time point of complete media exchange depends on the chosen retention time and needs to be taken into account for downstream analyses.

• 62.On the last day of continuous cultivation, collect as much sample volume from each vessel as desired before preparing the systems for decontamination and disassembly.

Decontamination and disassembly of ex vivo cultivation systems

Timing: 2–3 days, working time: 4 h per cultivation system

After the last sample was collected from the cultivation vessels, the cultivation systems need to be decontaminated. This is followed by the disassembly and cleaning of the cultivation vessels.

• 63.Ensure that the bioreactor is stopped (using eve® software or the touch screen).
• 64.
  Prepare the bioreactors for decontamination as follows:

  Note: Cleaning is facilitated by removing the cultivation media from the bioreactors and autoclaving them with ddH₂O.

  • a.
    Remove acid, base and media from the silicone hoses connecting the reagent bottles with the adaptation ports by pressing the respective rocker switches to the left. Place clamps close to the hose ends, disconnect the hoses from the reagent bottles and cover the hose ends with aluminum foil.

    Note: Ensure that the pump heads do not come in contact with acid or base to avoid material damage.

  • b.
    Place a clamp close to the end of the silicone hose connected to the waste pipe. Remove the hose from the waste bottle and cover the end with aluminum foil.
  • c.
    Disconnect pH and redox sensors, spargers and exit gas coolers from the basic unit.
  • d.
    Remove the heat sensors from the immersion pockets.
  • e.
    Place a clamp on the silicone hose of the dip tube and remove the Super Safe Sampler from the hose.

    • i.
      Remove the filter including silicone hose and syringe from the Super Safe Sampler.
    • ii.
      Place the Super Safe Samplers in a bottle filled with ddH₂O. Autoclave 121°C, 15 min.
  • f.
    Remove the clamps from the silicone hose of the exit gas coolers.
  • g.
    Remove the mounting plate with pump heads from the drive shafts on the basic unit. Fit it into the vessel holder.
  • h.
    Tighten the bracket between the thermal blocks on the basic unit to release the vessel holder.
• 65.Shut down the system using the touch screen and switch off the device with the power switch on the basic unit.
• 66.Autoclave the cultivation system: 121°C, 15 min

CRITICAL: Ensure that the electronic ends of sensors are not in contact with aluminum foil or metal clamps and that the clamps were removed from the silicone hoses of the exit gas coolers.

• 67.Disassemble, clean and store all parts according to the manufacturer’s instructions.

Note: If the system is used regularly, it does not need to be disassembled completely. Accessories like bearing holders, addition port adapters, hose holders, stirrer shafts, baffles, impellers, flow deflectors, magnetic couplings and immersion pockets for temperature sensors can stay mounted to the vessel top plates (be careful when laying the top plates onto a flat surface to not damage any accessories).

Note: The cleaning is easier when done immediately after the experiment ended.

CRITICAL: Ensure that the centering bearing remains in the triangular plate during cleaning. Consider pouring vessel contents through sieve to minimize the risk of losing it.

gDNA isolation, 16S rRNA amplicon sequencing and circadian analysis

Timing: multiple weeks

After the ex vivo cultivation is finished, the samples need to be prepared for 16S rRNA amplicon sequencing and subsequent circadian analysis. Circadian cosinor-fitting for ex vivo samples is performed using an in-house code by Isaiah Ting based on van der Veen and Gerkema¹⁵ using GraphPad Prism.

Note: Circadian cosinor-fitting can be performed using any graphical program; it does not require GraphPad Prism.

• 68.gDNA isolation, sequencing of the 16S rRNA amplicon and data analysis using the IMNGS platform and the R pipeline Rhea is performed according to an updated version of Reitmeier et al.¹⁶ with IMNGS2.
• 69.
  The circadian analysis is carried out as follows:

  • a.
    Organize the data in two columns: A) time and B) measure.

    Note: Data should cover at least one full cycle (i.e., 24 h), but ideally multiple cycles.

  • b.
    In GraphPad Prism, choose XY-Table to fit a user-defined, non-linear function using the Analyze buttton followed by generating a “NEW” equation.
  • c.
    Enter the following function for cosinor fit with a sloping mesor:

    $$Y=M+S\times X+A\times \cos \left(\frac{2\times \pi \times \left(X-\Phi \right)}{\tau }\right)$$
    M = Mesor; S = Slope; A = Amplitude; $\Phi$ = Acrophase; $\tau$ = Period

    Note: Fitting can be improved by setting initial values for M = mean of all data (∗YMID), A = Maximum value – Minimum value (∗YMAX-YMIN), $\tau$ = 24, $\Phi$ = Time of maximum value (Value of X at YMAX). If desired, bounds can also be set for each parameter, including i.e., enforcing a positive amplitude (A>0, Costrains Tab) (see Figure 6).

  • d.
    To statistically test the alternative hypothesis of a significant cosinor fit, choose the “Compare” tab to compare two equations using the Extra sum-of-squares F test.

    Note: This will contrast the cosinor function to the Null hypothesis reduced model comprising of a sloping straight line (i.e., with the following function).

    $$Y=M+S\times X$$

    Note: For ease of use the generic straight line in GraphPad Prism can be used (see Figure 7).

  • e.
    If the statistical comparison supports the alternative hypothesis that the data is best described by a cosinor wave, ensure that the confidence interval of the amplitude does not include 0 (zero), in which case the function is not periodic (see Figures 8 and 9).

Figure 6.

Screenshot of setting initial values for a new equation as described in step 69.c

Figure 7.

Screenshot of hypothesis testing as described in step 69.d

Figure 8.

Screenshots of outputs generated in step 69.e

Figure 9.

Cosinor fits of data from Vessel A and B for which output data are shown in Figure 8

Calculation of transfer efficiency

Timing: 2 h

Following the 16S rRNA amplicon sequencing and data analysis, the proportion of zOTUs transferred from the donor sample into the cultivation vessels can be calculated.

• 70.Use values given in “OTUs_Table-norm-rel” from Rhea script “1. Normalization”. Copy values for the donor sample, all inocula and day 7 of media 1 in all cultivation vessels into an empty Excel sheet.
• 71.Calculate means for the inocula (“Inocula-Mean”) as well as the duplicate cultivation vessels (“Vessel-Mean”).
• 72.Sort zOTUs according to abundance in the human donor sample (high to low).
• 73.Calculate the sum of all zOTUs in the donor sample, Inocula-Mean and Vessel-Means.
• 74.Count the zOTUs present in the donor sample and Inocula-Mean (“Donor+Inocula only zOTUs”).
• 75.Count the number of “Donor+Inocula only zOTUs” in the Vessel-Means.
• 76.Calculate transfer efficiency for each duplicate as follows:

$$Transferefficiency(\%)=\frac{A}{B}\times 100$$

A = “Donor+Inocula only zOTUs” in Vessel-Mean; B = zOTUs in donor sample + Inocula-Mean.

• 77.Visualize the results using a display program (i.e., GraphPad Prism).

Expected outcomes

This protocol uses ex vivo cultivation to investigate the effect of dietary components on the intestinal microbiota and its metabolites. To this end, human stool samples are used to inoculate the cultivation vessels and dietary components are integrated into cultivation media.

Experiments are carried out in duplicates, which is why it is important to check the similarity of the duplicate vessels regarding α-diversity and abundance of different taxa over time. Take into account any issues that arouse over the course of the experiment in the respective vessels to evaluate differences between cultivation vessels.

In our experience, the microbial communities adapt during continuous culture resulting in stable α-diversity and relative abundances from day 7 onwards. To judge the microbial transfer from the donor stool sample into the cultivation vessels, the transfer efficiency can be calculated (see major steps 70–77). This takes into account the zOTUs present in the donor sample and inocula compared to the zOTUs present in the cultivation vessels on day 7 of continuous culture (see Figure 10). Furthermore, α-diversity and microbial abundances can be compared between donor sample, inocula and continuous culture.¹ Using the R pipeline Rhea, β-diversity can be calculated between these groups and visualized in MDS or NMDS plots.¹

Figure 10.

Efficiency of transfer of zOTUs from infant donor samples into ex vivo cultivation system after 7 days of continuous culture with one inoculation

Transfer efficiency represented as mean of two technical replicates. n = 2.

After seven days of continuous culture when the microbial community composition stabilized, the impact of a dietary component and/or the presence of rhythmic taxa can be investigated. The former can be done by comparing α-diversity and relative abundances from media 1 vs. media 2 in each cultivation vessel as well as computing β-diversity for these groups. The same group comparisons can be applied to assess the effect on metabolite production.

The presence of rhythmic taxa can be determined by applying the above mentioned script.

Limitations

We established a protocol to evaluate the influence of different dietary components on the human intestinal microbiota and its metabolites in an ex vivo setting. Even though we aim to mimic human colonic conditions, due to limited access to human intestinal content, microbial communities for inoculation were derived from fecal samples. These samples were collected by study participants in an aerobic environment and processed by study staff as quickly as possible. Nevertheless, the exposure to oxygen potentially decreased the abundance of anaerobic species.

One important cornerstone of the transferability of results generated by these experiments to the human in vivo situation is the proportion of bacterial species that can be transferred from the fecal material into the bioreactor (“transfer efficiency”). Due to human variability, it is impossible to predict which bacterial species will be transferred into the bioreactor. We were able to increase the transfer efficiency to 53% in case of adult fecal material by inoculating twice instead of once. In case of infant fecal material, the total number of bacterial species in the donor sample is usually lower compared to the adult situation but more of those species can be transferred into the bioreactor, with transfer efficiencies around 80% under the indicated conditions.

The expansion of microbial communities from the same donor sample can vary between bioreactors, resulting in a vessel effect. To minimize this effect, it is important to apply the same settings (pH, temperature, feed rate, etc.) to all bioreactors. Further, it is advised to perform technical duplicates for each donor sample and condition (i.e., cultivation media mimicking a certain diet).

It is important to invest time into designing the experimental media and selecting suitable donor samples before the start of the experiment as both majorly affect the ability to address a certain research question. Digestibility indices can be applied to approximate the amount of target ingredients reaching the colon in vivo. Keep in mind that these indices are approximations and may not be available for every ingredient. Of similar importance is to plan downstream analyses before the start of the experiment to collect enough sample material and to store it appropriately (i.e., in stabilizers for long-term storage).

The investigation of bacterial rhythmicity requires the collection of samples from the cultivation system at least every 4 h over two days. The analysis can be improved by increased sample collection (i.e., hourly).

Frequent sample collection from the cultivation system, preparation of cultivation media and monitoring of the experimental conditions result in a labor-intense and expensive experimental setup. Hence, this protocol is suitable to answer direct questions with thought out experimental designs but not to test an infinite number of donor samples or cultivation media.

Troubleshooting

Problem 1

The stirrer shaft tip is no longer located in the centering bearing (major step 9).

Potential solution

Disconnect both cultivation vessels of the affected system from the basic unit. Remove temperature sensors and sensor cables from pH and redox sensors. Place 2 clamps on the silicone hose of the gas entry between the sparger and the air filter and remove the plug from the N₂ supply. Remove the mounting plate with pump heads from the drive shafts and fit it into the vessel holder. Disconnect the exit gas cooler from the basic unit and tighten the bracket between the thermal blocks on the basic unit to release the vessel holder. Transport the vessel holder, all reagent bottles and the waste bottle to a sterile environment (i.e., a laminar flow hood) and place the vessel holder inside. Remove the clamping ring from the vessel top plate of the affected cultivation vessel. Lift up the vessel top plate as little as possible to re-insert the stirrer shaft tip into the centering bearing. Fasten the clamping ring around the vessel top plate and transport everything back to the basic unit. Reconnect the cultivation vessels to the basic unit (major steps 20–24, 26).

Problem 2

The touch screen does not give the option to calibrate a pH probe for one vessel (major step 13).

Potential solution

On the touch screen, select the correct vessel and open the “Controller” menu. Click on “pH” and open the “Options” menu. Under “Misc”, ensure that the option “can be calibrated” is selected.

Problem 3

There was a problem (i.e., silicone tube burst, air filter fell out of exit gas cooler) during autoclaving of the cultivation system before the sterility was tested (major step 19).

Potential solution

Replace any silicone tube that burst during autoclaving (ensure to use the right diameter), secure the air filter back into the exit gas cooler and restore the condition the cultivation system had before autoclaving. Remove any clamps you might have placed on the silicone hose on the filters of the exit gas coolers. Autoclave the system again: 121°C, 15 min.

Problem 4

There is rust on the magnetic coupling after autoclaving of the cultivation system before the sterility was tested (major step 19).

Potential solution

Remove any ddH₂O in the cultivation vessels and store the pH and redox sensors according to the manufacturer’s instructions. Add 5% citric acid to the cultivation vessel until the magnetic coupling is covered. Incubate for at least 16 h (i.e., overnight) (refer to manufacturer’s instructions for more details). If the rust is gone, rinse the vessel and accessories thoroughly with ddH₂O and prepare the system for autoclaving as before (major steps 10–18). If the rust is not gone, inform the technician responsible for you.

Problem 5

The air filter fell out of the silicone hose on top of a reagent bottle (media, acid or base) or waste bottle while removing the aluminum foil (major step 25).

Potential solution

Cover the silicone hose with the inside of the aluminum foil you removed. Insert a sterile air filter (pore size 0.2 μm) into the hose.

Problem 6

No bubbles are visible in the cultivation media (major step 26).

Potential solution

The flow rate depends on the gas pressure in the facility. Find the right settings for your facility by increasing the N₂ influx using the touch screen until bubbles are visible. Note that depending on the location of the sparger, bubbles can be more or less visible in one vessel. Ensure that the amount of bubbles is similar in all cultivation vessels. Adjust the N₂ influx of the other cultivation vessels if necessary. Use the determined set point for following experiments and check it regularly.

Problem 7

The temperature inside the vessel is not changing but the temperature control is turned on (major step 27).

Potential solution

Ensure that the temperature sensor is immersed into the pocket for the temperature sensor until it hits the bottom.

Problem 8

The stirrer is not moving even though it was turned on using the touch screen (major step 27).

Potential solution

Ensure that you turned on the stirrer for the correct cultivation vessel. Often, the stirrers only start moving after a minute if set to 40 rpm. To shorten the time, consider increasing the speed to 100 rpm until it starts to move and then reduce it to 40 rpm. In case of inconsistent stirring speed or if the stirrer does not start to move even after a few minutes, contact the technician responsible for you.

Problem 9

The pH is not changing upon addition of acid or base (major step 28).

Potential solution

Ensure that the pH probe is connected correctly to the basic unit and that the stirrer is turned on. If necessary, disconnect and slowly reconnect the sensor cable. Constantly check the touch screen for changes in pH output. Add 5 more drops of acid or base and check for changes in pH.

Problem 10

It is not possible to draw up cultivation media using the super safe sampler or air is drawn into the Luer Lock syringe (major step 39).

Potential solution

When no media is drawn up, check that you removed the clamp from the silicone hose connecting the dip tube to the super safe sampler. If air is drawn into the syringe, check that all parts of the super safe sampler are connected tightly. Carefully tighten them if necessary. If it is still not possible to collect a sample with the super safe sampler: Use an inoculation needle to collect a sample. To do so, connect a syringe to a sterile inoculation needle, pierce the septum membrane and lower the needle into the cultivation media. Draw up the desired sample volume and remove the needle from the cultivation vessel. Proceed with aliquoting the sample in a sterile environment as described in major step 41.

Problem 11

It is not possible to rinse the dip tube with clean air because air cannot be pushed into the super safe sampler (major step 39).

Potential solution

Check if the air filter on the super safe sampler is blocked. In that case, remove the blocked filter and immediately replace it with a new sterile filter (pore size 0.2 μm). Add silicone hose (Ø = 2 mm) to the end of the filter and insert the syringe filled with air.

Problem 12

The media volume in one cultivation vessel is too low or there is a noticeable difference in the volume in two media bottles feeding into the cultivation systems with the same feed rate (major step 43).

Potential solution

• •No media is being fed into the system: Check that the media bottle is not empty, the feed is turned on and that the feed pump is working. Look into the cultivation vessel and watch out for drops of media falling from the addition port into the vessel. If no drops are visible, check the silicone hose and the small tube in the pump heads for blockages. Massage the tubes to remove blockages. Pay attention to not disconnect the hose connectors between the silicone hose and the small tube. In case this problem appeared after connecting a fresh bottle of cold media, consider warming up the tubes (i.e., warm water, hair dryer).
• •There is foam on the surface of the cultivation media: Stop the N₂ influx for 30 s using the touch screen and check if the foam is dissolving. Do not forget to turn the N₂ influx back on after the foam dissolved. Consider lowering the N₂ influx to the lowest possible value that still results in bubbles in the vessel and lowering the speed of the stirrer. Recapitulate if you added enough Antifoam B solution in the cultivation media before autoclaving.

Problem 13

The media volume in one cultivation vessel is too high or there is a noticeable difference in the volume of two waste bottles connected to cultivation systems that operate on the same feed rate (major step 43).

Potential solution

Ensure that the height of the waste pipe is correct. Increase the N₂ influx to 1% using the touch screen. Check the silicone hose connecting waste pipe and waste bottle for liquid accumulation and blockages. If you find a blockage, massage the hose until the blockage dissolves. If the entire hose is filled with liquid, it is possible that the adaptation port of the waste bottle is blocked. Place a clamp close to the end of the hose and change the hose to the spare addition port of the waste bottle. Remove the clamp. After the media volume returned to the desired value, reduce the N₂ influx to the standard setting using the touch screen.

Problem 14

The pH of the cultivation media is not correct (major step 43).

Potential solution

Ensure that the sensor cable is connected correctly and the bottles containing acid and base are not empty. If the pH is constantly 0.1 higher than the set point, lower the set point by 0.1.

Problem 15

It is not possible to inject the inoculum into the cultivation vessel using an inoculation needle (major step 47).

Potential solution

It is possible that the inoculation needle is blocked. Remove the blocked inoculation needle from the cultivation vessel and disconnect the syringe containing the inoculum with the needle pointing upwards. Immediately connect a new inoculation needle and continue with the inoculation as planned. After finishing the inoculation, clean the blocked needle thoroughly using the supplied wires for cleaning. Use a sonication bath if necessary. Check that the blockage is removed before wrapping the needle in aluminum foil and sterilizing at 180°C for 3 h. To avoid a shortage of inoculation needles, always prepare 2 per cultivation vessel.

Resource availability

Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Prof. Dr. Dirk Haller, (dirk.haller@tum.de).

Technical contact

Questions about the technical specifics of performing the protocol should be directed to and will be answered by the technical contact, Prof. Dr. Dirk Haller, (dirk.haller@tum.de).

Materials availability

This study did not generate any unique materials or reagents.

Data and code availability

Sequencing data reported in this protocol will be shared by the lead contact upon request. This study did not generate original code.

Acknowledgments

The Infantbio-II study was initiated and financed by Töpfer GmbH (Dietmannsried, Germany). The BISS-study was financed by the German Federal Ministry for Economic Affairs and Climate Action. Funding was received from the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation), project number 395357507 (SFB 1371, Microbiome Signatures). The ZIEL Institute for Food & Health of the Technical University of Munich provided technical support to perform the 16S rRNA gene amplicon analysis and the intervention studies. The graphical abstract was created using BioRender.com. We thank Jiatong Nie, Shengcheng Ren, Madeleine Spohn, Michael Vig Merino, and Isabel Zurblihn for their help in conducting the chemostat experiments. We thank Marjolein Heddes for her help to perform the circadian analysis. We thank the participants of both intervention trials for their time and involvement in our studies.

Author contributions

H.H. and D. Häcker performed the chemostat experiments; H.O. supervised the experimental procedures; S.K. and D.R.v.d.V. contributed to the circadian analysis; D. Haller evaluated and interpreted the data; H.H. wrote the manuscript draft; and all other authors critically evaluated the provided input, critically revised, and approved the final version of the manuscript.

Declaration of interests

The authors declare no competing interests.