Ventral stereotaxic coordinates have to be recalculated with respect to the open length of the cOFM guide and the spatial distribution (size and form) of the region of interest. The shaft length has to be chosen in a way that the open length is inside or immediately adjacent to the region of interest after implantation of cOFM guide.
Contact Joanneum Research for custom-made probes. Please note that customized probes may differ in price and delivery time. Many brain regions can be addressed by standard probes. However, if a custom probe is needed, they should follow the cOFM scheme:
Y should always be 1 mm or 2 mm.
X should be between 1 mm and 20 mm +/- 0.2 mm
Probe performance is guaranteed only for standard probes.
Unlike microdialysis, OFM has no membrane and therefore, no membrane swelling step is necessary. However, we suggest to perform a run-in phase at the sampling flow rate for at least one hour before starting sampling to achieve a stable flow rate (refer to the Instructions of Use for the cOFM probe).
Take care that the sampling insert and tubing are free of air bubbles before inserting them into the guide.
No, cOFM probes are gamma irradiated before delivery and autoclavation is not recommended. cOFM probes might be destroyed during autoclavation due to high temperatures.
No, there is no cap for the healing dummy.
For implantation, animals have to be anesthetized with appropriate anesthetics (e.g. isoflurane or ketamine) for 30 to 45 min. Then let the BBB recover for 14 days.
To replace the Healing Dummy with the Sampling Insert animals have to be sedated again. After removing the Lock, the Healing Dummy can be carefully removed from the OFM guide and replaced by pre-flushed Sampling Insert. Afterwards secure it with the Lock. Taking account the size of the pieces, Joanneum Research recommends the use of tweezers. Also, refer to the user manual of the cOFM probe, that can be downloaded from www.openflowmicroperfusion.com/info.
No, cleaning during the healing process is not necessary. We recommend keeping the healing dummy in place during the whole healing process until it is exchange with the sampling insert. Removing the healing dummy during the healing process may cause additional or new injuries and thus prolong the healing process.
No, cOFM probes are manufactured of PEEK and are not containing any metal, and can be scanned using MRI. But take into consideration that probes are attached to the scull using metal screws.
Yes, the OFM pump (MPP102PC) has push and pull options in the same unit. Either one pump head can be used for push and the second pump head for pull or, with special tubing (e.g. OFM-PP1-LT-1000), push and pull modes can be performed using only one pump head.
Calibration is necessary as different tubing materials and diameters are available. The calibration factors are pre-determined to save you time and money and improve your research quality. The calibration factor are stated on the label of the packaging. You can find more details in the MPP102PC user manual.
Flow rates can be adjusted between 0.1 – 10 µl/min with an accuracy of less than ± 20%. We recommend using pre-calibrated tubing to ensure sufficient accuracy. Furthermore, we suggest having the same flow rate for PUSH and PULL to avoid altering the pressure within the target tissue of the experimental animal.
Lowest flow rate: 0.1 µl/min
Highest flow rate: 10 µl/min.
Flow rates can be adjusted between 0.1 – 10 µl/min with an accuracy of less than ± 20%.
We recommend using pre-calibrated tubing to ensure sufficient accuracy.
Yes, we recommend using the central channel to ensure high sampling quality if you are not using all three channels. If a perfusate bag is used, unused channels need to be pinched off directly after the perfusate bag luer secured connector by pinch clamps. To reduce user errors, we recommend the use of single channel tubing to operate one single probe and the use of three channel tubing to operate up to three probes.
No, the battery does not recharge. But if the pump is operated on power supply the battery will not be discharged because it serves as a buffer battery in case the power supply fails. If the pump is not used for more than one week, remove the battery to avoid damaging the pump by battery leakage.No, the battery does not recharge. But if the pump is operated on power supply the battery will not be discharged because it serves as a buffer battery in case the power supply fails. If the pump is not used for more than one week, remove the battery to avoid damaging the pump by battery leakage.
JOANNEUM RESEARCH is considering a CPU control for the next version but it is not yet available for the current version.
There are tubing available from JOANNEUM RESEARCH that support operating up to 3 probes in parallel (e.g. OFM-PS3-75, Three Channel Push Tubing or OFM-PL3-75 Three Channel Pull Tubing). Please refer to the OFM pump user manual, which is available on the JOANNEUM RESEARCH webpage.
In theory, using three tubings at one pump head allows you to use three separate animals, as long as you are able to achieve a stable sampling process. However, an excessive tubing length might be counterproductive to achieve stable sampling. Therefore, Joanneum Research suggests one pump for three channels only for ex vivo experiments, whereas for animal studies we suggest using one pump for each animal.
Flush mode is generally used to flush the tubing and get rid of air bubbles. Thus, the pump is operated at a freely selectable flowrate of up to 10 µl/min. The flush mode allows you to select the amount of time and the flowrate at which the system will be flushed and what happens after the time has elapsed e.g. "Auto Pump", "Confirm Pump, "Auto Stop", "Confirm Stop". Please refer to the OFM pump user manual available on www.openflowmicroperfusion.com/info.
The second flush is identical to first flush mode, but allows you to set different parameters. This might be necessary for some experiments.
In general, we recommend to flush the tubing until it is air bubble free and then to perfuse it at the desired flow rate of e.g. 0.3 to 1.0 µl/min.
cOFM probes are normally used at the same flow rate for P1 and P2 (e.g. 0.3 to 1 µl/min). However, as the pump is also designed to operate dermal probes, flow rates might vary depending on the experiment. Joanneum Research recommends operating dermal probes
(a/d OFM-P-15) with a slightly higher PUSH- than PULL-flow rate e.g. 1.1/1.0 µl/min.
Tubing are precalibrated, thus the flow rate shown on the pumps display reflects the true flow rate if the “Tubing Adj %” is set to the values shown on the packaging.
Yes, pump can work with just a battery.
Typical operating time (with a new battery) is up to 48 hours at 1.0 μl/min and up to 24 hours at 10 μl/min. For longer operating time, change of battery is necessary or operation with power supply. Please refer to the OFM pump user manual available on www.openflowmicroperfusion.com/info.
Volume loss is possible if the push rate is higher than the pull rate. For proper cOFM sampling, we highly recommends the use of an OFM pump, which has guaranteed flow rate accuracy. Alternatively, syringe pumps can be used, as they have a comparably accurate flow rate over time.
For better sampling results, we recommend using the OFM perfusate bag, which is delivered fully sealed in gamma irradiated protective packaging. However, if necessary, you can use any container filled with any perfusate.
The perfusate composition is crucial for successful cOFM sampling. We suggest performing an in vitroadsorption test prior to the in vivo sampling to assess the degree of unspecific adsorption of the substance of interest to the cOFM system.
Please refer to the user manual of the perfusate bag. It shows in a step-by-step manner how to install and fill the perfusate bag in an air bubble free way.
1. Unpack tubing and perfusate bag.
2. Remove red protection cap of perfusate bag by turning it counter clockwise.
DO NOT remove needleless injection port.
3. Connect perfusate bag with the push tubing by turning the luer-connector clockwise.
4. Fill the perfusate bag with a (sterile) syringe via the needleless injection port.
CAUTION: DO NOT use aggressive fluids (alcohols, acids, bleaches, etc.) or hot fluids
(> 45°C) as perfusate (see instructions for use of perfusate bag).
With single channel tubing, one single probe can be operated. Three channel tubing allow operating up to 3 probes simultaneously.
No, other tubing than the ones developed for OFM will damage the OFM pump.
No, the tubing does not have to be switched after every use.
The tubing are gamma irradiated, but biofouling can still occur after using the tubing for a prolonged period of time, especially under unsterile conditions. Furthermore, tubing wear out over time, but can be typically used for 72 hours. If tubing are used longer, check the flow rate by measuring sampling time and volume and calculate the actual flow rate (flow (µl/min) = volume (µl) / time (min). If the tubing is not used for a couple of days between two sampling sessions, open the pump head and remove it from the pump. If multiple sampling sessions in experimental animals are performed, we recommend one day of rest for the experimental animal to recover. For every sampling session, use a lock to secure the sampling insert and use new tubing systems to guarantee the best sampling quality.
We recommend using a scalpel or a brain matrix blade with a 45° angle.
No, the low bind tubing OFM-T1-200-LB and OFM-T2-100-LB come in standard length and can be shortened depending on your needs.
Air bubbles can enter the tubing system due to the following three reasons:
(Also refer to the following picture)
1.) The Guide was not properly attached to the cranial bone of the animal or was loosened during manipulation (e.g. while inserting the Sampling Insert). In this case, re-implantation is necessary.
2.) The Sampling Insert leaks. In this case, check if the gasket is in place.
3.) Air bubbles enter between tubing and Sampling insert. In this case, use a drop of cyan acrylic glue.
No, the cOFM sampling insert is not causing damage to the brain during sampling. But, the shaft of the cOFM guide causes a trauma and a disruption of the blood-brain barrier during implantation. After implantation, the healing dummy inside the cOFM guide prevents ingrowth of brain tissue during blood-brain barrier re-establishment and trauma abatement. For sampling the healing dummy is replaced by the sampling insert. Unlike in microdialysis, where the membrane is longer than the guide (stylet) and might cause additional damage during insertion, the cOFM sampling insert is shorter than the guide and thus causes no addional damage (it is moved inside a cage).
Yes, cOFM allows sampling of hydrophilic to lipophilic substance regardless of molecular size. However, the whole setup (open length, flow rate, perfusion fluid, and additional materials in the system like connectors) has to be adapted to achieve optimal results.
cOFM can be calibrated (= recovery estimation) for absolute quantification of substances in the brain. Influencing factors are the size of open length, flow rate, and perfusate composition. Additionally, the physical characteristics of the substance of interest can influence the recovery factor (e.g. adsorption to components of the cOFM system). ISF that is sampled by cOFM is diluted, but the overall ISF composition is left unchanged.
The in vitro recovery of cOFM assesses the adsorption of the substance of interest to the cOFM system. In vitro recovery tests should be done with the same system components that are used in the in vivo setup (cOFM probe type, tubing, etc.). Once the adsorption is assessed, in vitro recovery can be used as a correction factor for in vivo results. In vitro recovery cannot be used as an in vivo recovery surrogate.
In vivo recovery is empirically assessed. Commonly used protocols can be found in the literature (e.g. no net flux method, retrodialysis etc.). The suited in vivo cOFM probe calibration method should be chosen according to the needs of the experiment.
The cOFM sampling insert can be replaced with the healing dummy after sampling, if you are not continuously perfusing the cOFM system. Leaving a sampling insert in the guide without continuous pumping may lead to clogging. cOFM sampling can be restarted on the next sampling day by following the protocol in the user manual. This procedure can be repeated as long as the cOFM probe and animal are in good condition.
We recommend one day of rest for the experimental animal to recover. For every sampling session, use new tubing systems to guarantee best sampling quality.
Typical continuous sampling sessions last from 24 h to 5 days. There is no evidence of tissue reactivity. We suggest changing the tubing system after 72 hours. If tubing are used longer, check the flow rate by measuring sampling time and volume and calculate the actual flow rate (flow (µl/min) = volume (µl) / time (min). For more information, please contact Joanneum Research
For OFM sampling, overnight flow rate stabilization is not required. You can insert the sampling insert in the morning. Afterwards we recommend to flush the tubing until it is free of air bubbles and afterwards to perfuse it with the desired flow rate of e.g. 0.3 to 1.0 µl/min.
We recommend replacing the healing dummy when it can be visually determined to be worn, but not necessarily between two sampling sessions. We also recommend using a new sampling insert for every sampling session.
After sampling the cOFM sampling insert is replaced by the healing dummy and animals can be returned to their home cages. Before insertion, the healing dummy should be cleaned with alcohol, followed by distilled water. When working with an anaesthetized animal setup, a minimum of one day of rest is recommended before the next sampling session. cOFM sampling can be restarted by following the protocol in the user manual. This procedure can be repeated as long as the cOFM probe and animal are in good condition.
The pump should be unplugged and stopped. We suggest using new tubing for each sampling session. The tubing are gamma irradiated, but still biofouling can occur after using the tubing for a certain period, especially under unsterile conditions. Furthermore, tubing wear out over time, but they can typically be used for 72 hours. If tubing are used for a longer time period, check the flow rate by measuring sampling time and volume and calculate the actual flow rate (flow (µl/min) = volume (µl) / time (min).
The healing dummy can replace the cOFM sampling insert after sampling and animals can be returned to their home cages. When working with an anaesthetized animal setup, a minimum of 1 day of rest is suggested before the next sampling session. cOFM sampling can be restarted by following the protocol in the user manual. This procedure can be repeated as long as the animal and the cOFM probe are in good condition.
The number of probes per animal depends on the brain region of interest and the size of the animal. When using more than one probe per animal, the surgeon has to consider the orientation of the lock, as it has to be removed when the healing dummy is replaced with the cOFM sampling insert.
Yes, JOANNEUM RESEARCH (JR) - HEALTH offers cOFM contract research services; based on broad experience, JR offers in vivo pre-clinical studies in animals (mouse, rat and pig) for brain, adipose and skin tissue. We perform metabolic tests to investigate the pharmacokinetics of active pharmaceutical ingredients when introduced enterally, parenterally or topically. We offer ex vivo studies in explanted human skin donated by individuals undergoing plastic surgery. We investigate the adsorption and liberation of topically active pharmaceutical ingredients. We function as One-Stop-Shop that offers:
JR - HEALTH is EN ISO 9001:2008, EN ISO 13485:2012 and GLP-Good Laboratory Practice certified.
Special hands-on training with OFM specialists from JOANNEUM RESEARCH is recommended to improve research quality but is not a prerequisite. All products come with a self-explaining user manual.
Following training programs are offered:
In addition, video conferencing and technical support or individual training sessions can be arranged.
User manuals are available for download from the JR webpage by scanning the QR-code or entering the link into a web browser, printed on the package of the OFM products.
Yes, during the implantation of the cOFM guide, the BBB is manually disrupted. Therefore, a trauma abatement and BBB re-establishment time of 14 days is recommended prior to sampling (Birngruber et al., 2013. Clin. Exp. Pharmacol. Physiol. 40, 864–71). After these 14 days, the BBB integrity of animals with implanted cOFM guide is fully reestablished and comparable to control animals without cOFM.
cOFM probes can also be used as a tool for infusion of substances. For simple infusion (push only), the infusion factor is 100%. If substance or molecules are perfused (push-pull), the factor is different and depends on flow rate, open length and substance characteristics (diffusion in tissue, clearance from tissue, etc.).
JOANNEUM RESEARCH began the development of OFM technology about 15 years ago. Due to the great success and the new demands for OFM products in clinical/ preclinical studies, JOANNEUM RESEARCH developed cOFM probes within the last few years for cerebral and neurological brain studies.