Source Paper
The Monoiodoacetate Model of Osteoarthritis Pain in the Mouse
Thomas Pitcher, João Sousa-Valente, Marzia Malcangio
Journal of Visualized Experiments • 2016
Source Paper
Thomas Pitcher, João Sousa-Valente, Marzia Malcangio
Journal of Visualized Experiments • 2016
A major symptom of patients with osteoarthritis (OA) is pain that is triggered by peripheral as well as central changes within the pain pathways. The current treatments for OA pain such as NSAIDS or opiates are neither sufficiently effective nor devoid of detrimental side effects. Animal models of OA are being developed to improve our understanding of OA-related pain mechanisms and define novel pharmacological targets for therapy. Currently available models of OA in rodents include surgical and chemical interventions into one knee joint. The monoiodoacetate (MIA) model has become a standard for modelling joint disruption in OA in both rats and mice. The model, which is easier to perform in the rat, involves injection of MIA into a knee joint that induces rapid pain-like responses in the ipsilateral limb, the level of which can be controlled by injection of different doses. Intra-articular injection of MIA disrupts chondrocyte glycolysis by inhibiting glyceraldehyde-3-phosphatase dehydrogenase and results in chondrocyte death, neovascularization, subchondral bone necrosis and collapse, as well as inflammation. The morphological changes of the articular cartilage and bone disruption are reflective of some aspects of patient pathology. Along with joint damage, MIA injection induces referred mechanical sensitivity in the ipsilateral hind paw and weight bearing deficits that are measurable and quantifiable. These behavioral changes resemble some of the symptoms reported by the patient population, thereby validating the MIA injection in the knee as a useful and relevant pre-clinical model of OA pain. The aim of this article is to describe the methodology of intra-articular injections of MIA and the behavioral recordings of the associated development of hypersensitivity with a mind to highlight the necessary steps to give consistent and reliable recordings.
Objective: To induce osteoarthritis-like pain and joint damage in mice through intra-articular monoiodoacetate injection and measure resulting mechanical hypersensitivity and weight bearing deficits
This is a Intra-articular Monoiodoacetate Injection protocol using mouse as the model organism. The procedure involves 20 procedural steps, 8 equipment items, 10 materials. Extracted from a 2016 paper published in Journal of Visualized Experiments.
Model and subjects
mouse • Not specified • unknown • 8-10 weeks • Not specified • 8
Study window
~10 week study window | ~17 hours hands-on
Core workflow
Animal housing and acclimatization • Prepare monoiodoacetate solution • Anesthetize mice
Primary readouts
Key equipment and reagents
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House 8-10 week-old mice in groups of 5 under a 12 hr light/dark cycle (lights on at 7:00 AM) with food and water ad libitum. Allow mice to acclimatize for 1 week prior to starting the experiment.
Note: Randomize and cage mice in groups of 5. Use animal numbers as codes to blind the experimenter to treatments. Use body weights as parameters for randomization.
“House 8-10 week-old mice in groups of 5 under a 12 hr light/dark cycle (lights on at 7:00 AM) with food and water ad libitum. Let the mice acclimatize for 1 week prior to starting the experiment.”
On the day of injection, freshly prepare the solution of monoiodoacetate in sterile saline (0.9% NaCl) at the desired concentrations. Prepare sterile saline for injections in a separate group of control mice.
Note: Highest recommended dose is 1 mg in 10 µl. Monoiodoacetate is very toxic - wear gloves and mask when handling powder and preparing solution. Solution should be sterile filtered with a 0.22 µm filter.
“On the day of injection, freshly prepare the solution of monoiodoacetate in sterile saline (0.9% NaCl) at the desired concentrations. The highest recommend dose of MIA is 1 mg in 10 µl. The solution should be sterile filtered with a 0.22 µm filter.”
Place mice in a chamber delivering 2% isoflurane in O2 mixture (flow rate 1.5 L/min). Transfer anesthetized mice to the nose cone section, which also delivers the 2% isoflurane-O2 mixture to maintain anesthesia during injection. Place vet ointment on the eyes to avoid drying out.
Note: Confirm anesthesia by checking the animal's lack of response to a pinch stimulus on the hind paws.
“Anesthetize mice using an anesthetic trolley by first placing them in a chamber delivering 2% isoflurane in O2 mixture (flow rate 1.5 L/min) and then transfer mice to the nose cone section, which also delivers the 2% isoflurane-O2 mixture, and as such, maintains anesthesia during injection.”
Once the animal is under anesthesia, place it on its back. Wear surgical gown, gloves, and mask while performing injection procedure.
Note: Confirm anesthesia by checking the animal's lack of response to a pinch stimulus on the hind paws.
“Once the animal is under anesthesia, place it on its back. Wear surgical gown, gloves, and mask while performing injection procedure.”
Trim and wipe the area surrounding the knee joint with alcohol. Povidone iodine or chlorhexidine can be used as alternative disinfectants.
Note: The patellar tendon (white line below the patella) will become visible after disinfection.
“Trim and wipe the area surrounding the knee joint with alcohol. Povidone iodine or chlorhexidine can be used as well for disinfection. The patellar tendon (white line bellow the patella) will become visible.”
To stabilize the injection site, keep the knee still in a bent position by placing the index finger beneath the knee joint and the thumb above the anterior surface of the ankle joint. Joint preference is not required.
Note: This positioning is critical for accurate injection.
“In order to stabilize the injection site, keep the knee still, in a bent position, by placing the index finger beneath the knee joint and the thumb above the anterior surface of the ankle joint.”
Run a 26 G needle attached to a syringe horizontally along the knee (so as not to pierce the skin with the tip) until it finds the gap beneath the patella. Apply gentle pressure to mark the area and then lift the needle and syringe vertically for the injection.
Note: This step identifies the correct anatomical location for intra-articular injection.
“To find the precise site of injection, run a 26 G needle attached to a syringe horizontally along the knee (so as not to pierce the skin with the tip) until it finds the gap beneath the patella. Apply gentle pressure to mark the area and then lift the needle and syringe vertically for the injection.”
Insert the needle in the marked area, through the patellar tendon, perpendicular to the tibia. No resistance should be felt. Use thumb as a guide and inject superficial to the site of entry. Inject the MIA solution (10 µl at desired concentration).
Note: Absence of resistance indicates correct intra-articular placement. Discard the needle immediately in the sharps bin after injection.
“Insert the needle in the marked area, through the patellar tendon, perpendicular to the tibia. No resistance should be felt. Use thumb as a guide and inject superficial to the site of entry.”
After injection, massage the knee to ensure even distribution of the solution.
Note: This step ensures uniform distribution of MIA throughout the joint space.
“After injection, massage the knee to ensure even distribution of the solution.”
Place mice back into a clean home cage on a heated mat and allow them to recover. Keep constant vigilance on the animals until they regain suitable consciousness, which is measured by them regaining sternal recumbency. Once animals are recovered, return to their cage.
Note: It is suggested for best practice and training purposes that a dye is used and immediate post-mortem dissection performed to confirm correct localization of injection.
“Place mice back into a clean home cage on a heated mat and allow them to recover. Keep constant vigilance on the animals until they regain suitable consciousness, which is measured by them regaining sternal recumbency.”
Bring mice to the behavioral room and let unrestrained animals acclimatize in acrylic cubicles (8 cm x 5 cm x 10 cm) atop a wire mesh grid. Train mice by handling and 2 hr habituation to the cubicles for two days prior to von Frey hair application in order to limit stress and ambulation during application of von Frey hairs.
Note: On test days, habituate animals to the cubicles for up to 60 min prior to testing. Wear gowns, gloves, and masks during all behavioral experiments.
“Train mice by handling and 2 hr habituation to the cubicles for two days prior to von Frey hair application in order to limit stress and ambulation during application of von Frey hairs. On test days, habituate animals to the cubicles for up to 60 min prior to testing.”
Assess mechanical thresholds of both hind paws before MIA injection as baseline values using the up-down method with von Frey hairs.
Note: This establishes baseline withdrawal thresholds for comparison with post-injection measurements.
“Following the procedure described above (2.2.1-2.2.4), assess mechanical thresholds of both hind paws before MIA injection as baseline values.”
Starting with a stimulus strength of 0.07 g, apply von Frey hairs according to the up-down method: mark as X a withdrawal response and O an absence of response. Apply in ascending order of force, up to 1 g (cut-off force), until a response is detected. Re-test the paw by repeating with the filament that exerts a force below the one that produced a withdrawal. Then, apply the remaining filaments sequentially, by descending force, until no withdrawal occurs. Re-apply filaments in ascending order until a response is observed. Continue until a sequence of six responses is obtained (e.g., OXOXOX).
Note: Hold each hair in place for 3 sec or until the paw is withdrawn, the latter defining a positive response. Use 0.008, 0.02, 0.04, 0.07, 0.16, 0.4, 0.6, and 1.0 g fibers during testing.
“Apply calibrated von Frey hairs to the plantar surface of the hind paw until the fiber bends. Hold each hair in place for 3 sec or until the paw is withdrawn, the latter defining a positive response. Starting with a stimulus strength of 0.07 g, apply hairs according to the up-down method: mark as X a withdrawal response and O an absence of response.”
Obtain the 'k' value by referring to tabular values. Express paw withdrawal values as 50% paw withdrawal thresholds in grams using the formula (10 [Xr + K δ])/10,000 where Xr = value of last von Frey filament used in the sequence (in log units), k = tabular value, and δ = mean difference in forces between fibers. Where no response is detected, use the maximal response of 1 g.
Note: This calculation converts the up-down method response pattern into a quantitative threshold value.
“obtain the 'k' value by referring to tabular values. Express paw withdrawal values as 50% paw withdrawal thresholds in grams. Use the formula (10 [Xr + K δ])/10,000 where Xr = value of last von Frey filament used in the sequence (in log units), k = tabular value, and δ = mean difference in forces between fibers.”
After injection, assess thresholds of the ipsilateral and contralateral paws at regular day intervals for several weeks after MIA to ascertain the development of mechanical allodynia. For example, assess at 0, 3, 5, 7, 10, 14, 21, and 28 days after MIA injection.
Note: Animals are considered allodynic when they display a response to 0.1 g or less. Normal responses fall within 0.6-1 g range.
“After injection, assess thresholds of the ipsilateral and contralateral paws at regular day intervals for several weeks after MIA to ascertain the development of mechanical allodynia. For example, we report thresholds measured 0, 3, 5, 7, 10, 14, 21, and 28 days after MIA injection.”
Train each mouse to walk into a Plexiglass chamber on the weight incapacitance tester apparatus and sit in the holding box. Place the mouse in front of the holding box, lift the entrance up 45°, and allow the mouse to walk in and close the box. Allow the animals to move freely until they adopt a sitting posture.
Note: This training guarantees that the animal is still and not leaning on either side of the chamber.
“Train each mouse to walk into a Plexiglass chamber on the apparatus and sit in the holding box. Place the mouse in front of the holding box, lift the entrance up 45°, and allow the mouse to walk in and close the box. This training takes at least two days and guarantees that the animal is still and not leaning on either side of the chamber.”
Calibrate the weight incapacitance tester before use with a 100 g check weight or according to equipment instruction.
Note: Proper calibration is essential for accurate weight bearing measurements.
“Calibrate the instrument before use with a 100 g check weight (or according to equipment instruction).”
Assess weight bearing changes before MIA injection as baseline values. Make sure that each hind paw is placed on the appropriate recording pad. The duration of each measurement takes 1 sec, as per the manufacturer's instructions. Collect three measurements of the weight borne on each hind paw from the recording pad for each recording session and use the mean value to calculate the difference in weight borne by ipsilateral and contralateral paws.
Note: Express values as the difference between contralateral and ipsilateral paws in grams.
“Make sure that each hind paw is placed on the appropriate recording pad. The duration of each measurement takes 1 sec, as per the manufacturer's instructions. Collect three measurements of the weight borne on each hind paw from the recording pad for each recording session and use the mean value to calculate the difference in weight borne by ipsilateral and contralateral paws.”
Repeat weight bearing assessments at regular intervals over several weeks to ascertain the development of gait changes. For example, assess at 0, 3, 5, 7, 10, 14, 21, and 28 days after MIA injection. Express values as the difference between contralateral and ipsilateral paws in grams.
Note: A normal weight bearing value of 50% represents an equal weight distribution across ipsilateral and contralateral hindlimb. Animals considered hypersensitive display a weight bearing change of approximately 45%.
“Then, repeat assessments at regular intervals over several weeks to ascertain the development of gate changes. For example, we report thresholds measured on 0, 3, 5, 7, 10, 14, 21, and 28 days after MIA injection.”
Measurements of mechanical thresholds and weight bearing deficits can be performed in the same mice, as neither end point affects the other.
Note: This allows for comprehensive assessment of pain-like behavior in individual animals.
“Measurements of mechanical thresholds and weight bearing deficits can be performed in the same mice, as neither end point affects the other.”
This section explains what the experiment is doing, which readouts matter, what the data artifacts usually look like, and how the analysis should flow from raw capture to reported result.
To induce osteoarthritis-like pain and joint damage in mice through intra-articular monoiodoacetate injection and measure resulting mechanical hypersensitivity and weight bearing deficits
Objective
To induce osteoarthritis-like pain and joint damage in mice through intra-articular monoiodoacetate injection and measure resulting mechanical hypersensitivity and weight bearing deficits
Subjects
From papermouse • Not specified • unknown • 8-10 weeks • Not specified
Sample count
From paper8
Cohort notes
From paperHoused in groups of 5 under 12 hr light/dark cycle (lights on at 7:00 AM) with food and water ad libitum.
Animal housing and acclimatization (1 week)
Prepare monoiodoacetate solution (Day of injection)
Anesthetize mice (Until anesthesia is confirmed)
Position mouse for injection (Immediate)
Mechanical withdrawal thresholds (50% paw withdrawal threshold in grams) measured using von Frey hairs
From paperTwo-way repeated measurements ANOVA followed by Student Newman-Keuls post hoc test.
Artifact type
Endpoint measurements summarized by group or timepoint
Comparison focus
Compare endpoint magnitude between groups, timepoints, or both
Development of mechanical allodynia (response to 0.1 g or less)
From paperTwo-way repeated measurements ANOVA followed by Student Newman-Keuls post hoc test.
Artifact type
Endpoint measurements summarized by group or timepoint
Comparison focus
Compare endpoint magnitude between groups, timepoints, or both
Weight bearing distribution between ipsilateral and contralateral hind paws (expressed as difference in grams)
From paperTwo-way repeated measurements ANOVA followed by Student Newman-Keuls post hoc test.
Artifact type
Endpoint measurements summarized by group or timepoint
Comparison focus
Compare endpoint magnitude between groups, timepoints, or both
Weight bearing asymmetry (normal = 50% equal distribution; hypersensitive = approximately 45%)
From paperTwo-way repeated measurements ANOVA followed by Student Newman-Keuls post hoc test.
Artifact type
Endpoint measurements summarized by group or timepoint
Comparison focus
Compare endpoint magnitude between groups, timepoints, or both
Mechanical withdrawal thresholds (50% paw withdrawal threshold in grams) measured using von Frey hairs
From paperRaw artifact
Per-sample or per-animal endpoint measurements collected during the experiment
Processed artifact
Structured table with cleaned measurements ready for comparison
Final reported form
Summary statistics and between-group or across-timepoint comparisons
Development of mechanical allodynia (response to 0.1 g or less)
From paperRaw artifact
Per-sample or per-animal endpoint measurements collected during the experiment
Processed artifact
Structured table with cleaned measurements ready for comparison
Final reported form
Summary statistics and between-group or across-timepoint comparisons
Weight bearing distribution between ipsilateral and contralateral hind paws (expressed as difference in grams)
From paperRaw artifact
Per-sample or per-animal endpoint measurements collected during the experiment
Processed artifact
Structured table with cleaned measurements ready for comparison
Final reported form
Summary statistics and between-group or across-timepoint comparisons
Weight bearing asymmetry (normal = 50% equal distribution; hypersensitive = approximately 45%)
From paperRaw artifact
Per-sample or per-animal endpoint measurements collected during the experiment
Processed artifact
Structured table with cleaned measurements ready for comparison
Final reported form
Summary statistics and between-group or across-timepoint comparisons
Acquisition
Collect raw experimental outputs with enough metadata to preserve sample identity, condition, and timing.
Preprocessing / cleaning
Two-way repeated measurements ANOVA followed by Student Newman-Keuls post hoc test.
Scoring or quantification
Quantify the primary readouts for this experiment: Mechanical withdrawal thresholds (50% paw withdrawal threshold in grams) measured using von Frey hairs; Development of mechanical allodynia (response to 0.1 g or less); Weight bearing distribution between ipsilateral and contralateral hind paws (expressed as difference in grams); Weight bearing asymmetry (normal = 50% equal distribution; hypersensitive = approximately 45%).
Statistical comparison
Statistical method not yet structured for this page.
Reporting output
Report representative outputs alongside summary comparisons for Mechanical withdrawal thresholds (50% paw withdrawal threshold in grams) measured using von Frey hairs, Development of mechanical allodynia (response to 0.1 g or less), Weight bearing distribution between ipsilateral and contralateral hind paws (expressed as difference in grams), Weight bearing asymmetry (normal = 50% equal distribution; hypersensitive = approximately 45%).
Source links and direct wording from the methods section for validation and deeper review.
Citation
Thomas Pitcher et al. (2016). The Monoiodoacetate Model of Osteoarthritis Pain in the Mouse. Journal of Visualized Experiments
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