In 2010 the ARRIVE guidelines (Animal Research: Reporting of In Vivo Experiments) were set up to improve reporting of animal research. In 2020 an update of the guidelines was published:

https://doi.org/10.1371/journal.pbio.3000410

The reorganised guidelines were presented in 2 sets, ”The ARRIVE Essential 10” and ”The Recommended Set“.

Here we show the realisation of the ARRIVE guidelines for the GMC.

These items are the basic minimum to be included in a manuscript. Without this information, readers and reviewers cannot assess the reliability of the findings.

For each experiment, provide brief details of study design including:

a. The groups being compared, including control groups. If no control group has been used, the rationale should be stated.

b. The experimental unit (e.g. a single animal, litter, or cage of animals).

a. We compare a mutant group with the respective control group for each sex (female and male): The control group consists of littermates of the experimental group.

b. The individual mouse was considered the experimental unit within the project.

a. Specify the exact number of experimental units allocated to each group, and the total number in each experiment. Also indicate the total number of animals used.

b. Explain how the sample size was decided. Provide details of any a priori sample size calculation, if done.

a) We used a sample size of n=15 mice per genotype and sex. Consequently, the total number of mice used in one experiment is N=60. Reasons for deviations of this number (e.g. dead animals) are documented.

b) The selected sample size of 15 mice per sex and genotype is sufficient to find a medium difference of one standard deviation with a power of 0.8 and alpha of 0.05. For some parameters (e.g. categorical, yes/no) even less animals are sufficient.

Sample size estimation is based on long-term experience in high-throughput phenotyping. Since we conduct a high throughput screening we apply no a priory sample size calculations in the format of power considerations.

In the sense of 3R, we do not include reserve animals. The estimated sample size is sufficient to generate a robust data basis, even in the rare case of shortfall. Further reserve animals are not applicable, since it is strongly recommended to only use animals with the same handling during the screen, to ensure comparability.

a. Describe any criteria used for including and excluding animals (or experimental units) during the experiment, and data points during the analysis. Specify if these criteria were established a priori. If no criteria were set, state this explicitly.

b. For each experimental group, report any animals, experimental units or data points not included in the analysis and explain why. If there were no exclusions, state so.

c. For each analysis, report the exact value of n in each experimental group.

a. In general, all mice with genotypes that are to be compared  (e.g. homozygous versus wild type control animals) that result

from for the purpose appropriate breedings (e.g. heterozygous x heterozygous to generate homozygous and wild type animals) are considered to reflect a representative sample of the population and will be taken for the experiment as obtained by the breeding. If we observe single individual animals that obviously deviate from the population, where it can be unambiguously excluded that this deviation is caused by the mutation that is to be analyzed (e.g. hermaphrodites, long teeth as result from dysgnatia, or mice with injuries from fights with cage mates), and where the inclusion of this animal would systematically confound the experiment, are excluded as an a priori criterion.

Before the start of each individual experiment within the phenotyping pipeline every animal is examined for its physical condition. In case that the condition of the animal does not allow that the experiment can be performed in a safe and harmless manner for the animal, the test will be skipped for this animal. Additional exclusion criteria as well as stop criteria that are specific for each test are defined in the SOP for each test. For example, digit abnormalities exclude mice from grip. strength measurements. A test will be completely stopped as soon as two mice of the test cohort show any problems with the test conditions (e.g. anesthesia incompatibility in the X-ray test).

b. In general, no data point will be excluded from the analysis, unless there is clear evidence of e.g. technical failure of the experimental machine. For many of the parameters (like blood parameters) we have pre-established threshold values as indicators for invalid measurements. If the data point reaches the criteria, we have to expect that there was an undetectable failure, and the data point is replaced by a status code (e.g. above upper limit).

c.

a. State whether randomization was used to allocate experimental units to control and treatment groups. If done, provide the method used to generate the randomization sequence.

b. Describe the strategy used to minimize potential confounders such as the order of treatments and measurements, or animal/cage location. If confounders were not controlled, state this explicitly.

b) We control for all known influencing factors: our mice are maintained under specific pathogen-free conditions with standardized diet, acclimatization and standardized cage enrichment. All mice tested are located in the same rack within the housing room.

Female and male mice are separated in the cages, genotypes are mixed. Cage sizes are between 2 and 5 animals usually. The maximum age difference between youngest and oldest animal is 7 days.

To minimize potential confounders during the measurement, we standardize external factors as much as possible. All mice of a project will usually be measured concurrently with the same machine and by the same technician.

When it comes to testing the mice, we ensure that selection of the mice is balanced, meaning, for example, that we measure control and mutants in parallel or alternately. In tests where the order of the mice has an influence on the results (e.g. earlier versus later sample taking), we create lists defining the order of the measured mice that are balanced for the experimental and the control group.

Blood samples are randomized by the robot (the robot includes a specific programing for randomization of the samples in the plate).

Describe who was aware of the group allocation at the different stages of the experiment (during the allocation, the conduct of the experiment, the outcome assessment, and the data analysis).

In most tests, blinding is not needed, as the experimenter has no influence on the results of these tests, since the results are directly recorded by the machine. In the tests, where the experimenter might have an influence on the measurements, the experimenter is blinded, and we have SOP's prescribing how this will be ensured for each single test separately. Blood samples are blinded (the robot works with barcodes, not IDs of the mice), preparation by automation. In addition, we record metadata for each data point and monitor the influence of these metadata over time.

In some cases, blinding is not possible, e.g. when the genotype is obvious by simply looking at the mice.

Data analysis is automated via standardized R-scripts.

a. Clearly define all outcome measures assessed (e.g. cell death, molecular markers, or behavioural changes). b. For hypothesis-testing studies, specify the primary outcome measure, i.e. the outcome measure that was used to determine the sample size.

a) We measure more than 500 parameters in all screens.

b) Not applicable: No hypothesis testing study (screening project)

a. Provide details of the statistical methods used for each analysis, including software used.

b. Describe any methods used to assess whether the data met the assumptions of the statistical approach, and what was done if the assumptions were not met.

a) Data is analyzed using automated R-scripts (software version 3.4.4   (2018-03-15) -- "Someone to Lean On").

Since we have phenotyping data of control animals for almost 20 years, we know the distribution of every single parameter. Based on this knowledge, we selected the best statistical test. We have a customized analysis for each parameter including also complex models (e.g. LME's, body-weight as additional covariate).

For the exact statistical methods used in each screen, please refer to table 6a)

b) Please see above. Distributions of the parameters are regularly checked. We implemented different quality control plots like QQ plots, histograms, KS test, effect sizes etc. in our automated R-scripts used to analyze the data.

a. Provide species-appropriate details of the animals used, including species, strain and sub strain, sex, age or developmental stage, and, if relevant, weight.

b. Provide further relevant information on the provenance of animals, health/immune status, genetic modification status, genotype, and any previous procedures.

a) The genetic background of the animals is project specific (at least 3 generations controlled breeding (e.g. backcrossing) after mutation generation). The used female and male mice usually start the phenotyping pipeline from the age of 9 weeks until 18 weeks. Wild-type animals have to be littermates of the mutant animals. The maximum age difference between youngest and oldest is 7 days.

b) We request a health report for the mouse line according to the FELASA guidelines before start of the project. The decision whether we can accept the mice in our facility is done by independent veterinarians. In case that the mice would not be allowed to enter our facility, they will not even be bred (3Rs principle). A second, more recent, health report is requested before sending the mice.

The collaboration partner generates (kind of mutation is project specific) and breeds the cohort for the phenotypic analysis and provides the requested animal number (15 males and 15 females (mutants) and 15 males and 15 females (wild type)). The animals have to be sent when they are 7 weeks old, with complete genotype information.

For each experimental group, including controls, describe the procedures in enough detail to allow others to replicate them, including:

a. What was done, how it was done and what was used.

b. When and how often.

c. Where (including detail of any acclimatization periods).

d. Why (provide rationale for procedures).

a)  Within the screening pipeline (Gailus-Durner et al., 2005; Fuchs et al., 2009, Gailus-Durner et al., 2009, Fuchs et al., 2011, Fuchs et al., 2018) all clinically relevant organ-systems of the mice are analyzed systematically covering a broad range of tests to identify novel phenotypes. Procedures of the tests are indicated in each test-SOP (including information about e.g. calibration or quality control steps). Methods of the standardized primary pipeline are published (see table below).

b) The pipeline usually starts with animals at 9 weeks of age. Tests, order of the tests and frequency is depicted in the table above and in the workflow of the screening pipeline (https://www.mouseclinic.de/research/gmc-pipelines/screening-pipeline/index.html)

c) The animals are sent by the collaboration partner who performed the breeding and are directly transferred to the dedicated housing room in our SPF facility. The mice have an acclimatization period of 2 weeks before the first measurement. We have a dedicated lab room for each test procedure. Mice are housed in a housing room close to the lab room, and are brought to the lab for the test. Test-specific acclimatization periods are defined in SOPs.

d) not applicable

For each experiment conducted, including independent replications, report:

a. Summary/descriptive statistics for each experimental group, with a measure of variability where applicable (e.g. mean and SD, or median and range).

b. If applicable, the effect size with a confidence interval.

a/b) For each parameter, we calculate measures like mean, standard-deviation, effect size and respectively median and IQR in our standardized automatic R-scripts.

These items complement the Essential 10 and add important context to the study. Reporting the items in both sets represents best practice.

Provide the name of the ethical review committee or equivalent that has approved the use of animals in this study, and any relevant license or protocol numbers (if applicable). If ethical approval was not sought or granted, provide a justification.

The GMC holds a general license to run phenotype assessments in mice and all tests are approved by the responsible authority of the district government of Upper Bavaria. Individual projects are registered and approved by the government approximately 4 weeks before start.

Provide details of housing and husbandry conditions, including any environmental enrichment.

Animal housing is performed in strict accordance with directive 2010/63/EU and the local government. All mice are housed in individually ventilated caging (IVC) systems (Sealsafe plus, GM 500, Tecniplast, Buggugiate, Italy) under specific pathogen-free conditions (GMC) with a maximum cage density of five adult mice per cage. IVC systems operate with positive pressure. All mice receive autoclaved wood chips (Lignocel select fine, J. Rettenmaier & Soehne GmbH, Rosenberg, Germany) and paper stripes (Arbocel crinclets natural Rettenmaier & Soehne GmbH) as bedding and nesting material, enrichment (red houses),irradiated standard diet for rodents (Altromin 1314, Altromin Spezialfutter GmbH, Lage, Germany) and sterile-filtered tap water ad libitum. Mice are transferred to new cages in laminar flow cage changing stations in weekly intervals. Light is adjusted to a 12h/12h light/dark cycle with a 10 min period of dimmed light to simulate sunrise/sunset; temperature and relative humidity are regulated to 22 ± 2° C and 55 ± 10 %. Access via air sluices is restricted by an electronic key system, and it is mandatory to wear personal protective equipment (autoclaved trousers and shirts, dedicated shoes, surgical masks, hair bonnets and gloves). Health monitoring is based on quarterly exhaust air dust PCR analysis for all FELASA-listed agents by a commercial diagnostic laboratory (Mähler M et al., 2014). All mice tested are located in the same room and rack.

Provide a statement describing if and where study data are available.

Raw data can be requested (the data will be transmitted via fixed data formats such as CSV/XML).

Upload data of single knockout lines to the IMPC webpage ( https://www.mousephenotype.org )- the data can be downloaded directly from the webpage (https://www.mousephenotype.org/understand/accessing-the-data/). 

The phenotyping results of the analyzed mouse lines can be uploaded to our Open Access GMC phenomap (https://www.mouseclinic.de/– see results).

Upload the data to a generic repository on demand.