Characterizing Baseline Fixed Charge Density in Human Cervical Intervertebral Discs

Abstract

This study provides quantification of fixed charge density in human cervical intervertebral discs. Fixed charge density, which occurs due to negatively charged proteoglycans in the extracellular matrix, is a key determinant of the intervertebral disc osmotic environment and swelling properties. While regional fixed charge density patterns have been characterized in lumbar discs, they remain unexplored in cervical discs. Using fresh-frozen cadaveric cervical discs from five donors, fixed charge density was measured using a two-point electrical conductivity method. Glycosaminoglycan content and porosity were also assessed. Fixed charge density (0.18 ± 0.1 mEq/g wet tissue) was highest in the cartilage endplate region and significantly greater than in that in the annulus fibrosus (p = 0.006). No significant difference in fixed charge density was observed between the nucleus pulposus and annulus fibrosus. Glycosaminoglycan content (40.3 ± 14.4 μg/mg wet tissue) showed a strong positive correlation with fixed charge density across regions (r = 0.65, p = 0.0047). Unlike lumbar discs, fixed charge density was found to be more homogeneous between the nucleus pulposus and annulus fibrosus regions. This result likely reflects adaptations for reduced tissue swelling in cervical discs to accommodate lower weight-bearing demands and increased flexibility. The elevated fixed charge density in the cervical endplates may protect the intervertebral disc-vertebral bone interface, potentially to avoid mechanical damage in a kinematically more mobile environment. These findings establish key benchmarks for understanding cervical disc electro-biomechanics and may inform other cervical disc tissue-characterization efforts.

Introduction

Intervertebral discs (IVDs) are among the largest avascular cartilaginous tissues in the human body, which facilitate load transmission between vertebrae and enable spinal movements such as flexion, extension, lateral bending, and torsion (Alonso and Hart, 2014; Cortes and Elliott, 2014). Unlike lumbar discs, which primarily resist axial loads from body weight, cervical IVDs are structurally adapted for greater kinematic mobility (Bogduk, 2016). Adaptations such as smaller disc heights and cranially accentuated saddle-shaped geometries enhance rotational freedom with bending (Busscher et al., 2010; Pooni et al., 1986; Bogduk and Mercer, 2000; Bogduk, 2016; Kumar and Pai, 2020). Additionally, zygapophysial joints contribute to load sharing and guide torsional movements, reducing mechanical stress on cervical discs (Bogduk, 2016; Cortes and Elliott, 2014). At the tissue level, cervical IVDs share key features with other spinal discs, including the annulus fibrosus (AF), which laterally confines the NP, and cartilage endplates (CEPs), which interface with adjacent vertebrae. Nutrient transport into these avascular tissues occurs primarily through solute diffusion across the AF and CEP regions (Nachemson et al., 1970; Urban et al., 2004; Huang et al., 2014).

In lumbar discs, biomechanical and nutrient transport behaviors are closely linked to extracellular matrix properties such as porosity, permeability, and fixed charge density (FCD) (Jackson et al., 2012; Cortes et al., 2014; Wu et al., 2016, 2017; Buchweitz et al., 2024). FCD occurs from the presence of negatively charged carboxyl and sulfate groups on glycosaminoglycans (GAGs) embedded in the IVD matrix, and plays a central role in maintaining the osmotic environment within the disc (Urban and Maroudas, 1979; Stevens et al., 1979). FCD is critical for establishing osmotic pressure gradients between the IVD matrix and surrounding interstitial fluid, which sustains tissue hydration and increases its stiffness, while also reducing permeability, helping to resist compressive loads (Cortes et al., 2014; DeLucca et al., 2016). Furthermore, by modulating the osmolarity of the extracellular matrix locally, FCD may influence cellular metabolic activity (Wuertz et al., 2007; Neidlinger-Wilke et al., 2012; Wu et al., 2017). Reductions in FCD, linked to diminished proteoglycan synthesis or increased degradation, are strongly associated with IVD degeneration, making FCD a key marker of disc health (Urban and Roberts, 2003).

Limited research on the biochemical composition of cervical IVDs suggests that their proteoglycan and water content distribution is more homogeneous between NP and AF regions, and that they exhibit greater collagen content compared to lumbar discs (Scott et al., 1994; Tomaszewski et al., 2015; Bostelmann et al., 2017) (Supp. Table 1). These findings suggest that the spatial FCD distribution in cervical discs may also be more homogeneous. While GAG content could be considered as a proxy for FCD, these measurements, which are reported on a dry weight basis, do not consider the effects of tissue hydration on IVD electro-mechanical behavior (Urban and Maroudas, 1979). Moreover, FCD itself is a critical parameter for computational modeling of IVD biomechanics, particularly for simulating swelling pressure effects (Iatridis et al., 2003; Cortes et al., 2014; Yao and Gu, 2004, 2006). While regional variations in FCD have been characterized in lumbar discs (Urban and Maroudas, 1979; Sivan et al., 2006; Wu et al., 2017), comparable data for cervical discs are lacking, leaving a gap in understanding their electro-mechanical environment.

Two-point electrical conductivity analysis provides a reliable and sensitive method for quantifying FCD in the IVD, with advantages of cost, safety, and accuracy over tracer cation assays, MRI imaging, and mechanical indentation (Maroudas and Thomas, 1970; Urban and Maroudas, 1979; Chen et al., 2003; Lu et al., 2004; Jackson et al., 2009; Wu et al., 2017). This approach has also been validated for thin IVD tissues like the CEP (Wu et al., 2017). Using this method, we hypothesized that cervical IVDs would exhibit a relatively more homogeneous FCD distribution between the NP and AF, consistent with the enhanced mobility of these discs. Additionally, we expected higher FCD in the CEP than in other disc tissues, aligned with its role as a transitional mechanical barrier tissue (Wu et al., 2017; Buchweitz et al., 2024). This study aimed to quantify the regional distribution of FCD in cervical IVDs, assess ion diffusivity, and correlate these measurements with biochemical composition (i.e., GAG content and tissue porosity). These findings are intended to provide foundational insights into the electro-mechanical environment of cervical discs and support future research on cervical IVD biomechanical modeling and regional tissue characterizations.

Methods

Sample preparation

Five cadaveric cervical spines (mean age: 59 ± 13 years, all male) were obtained from a tissue procurement organization (Science Care, Phoenix, AZ) with institution approval. The Thompson grading scale was used to screen the discs for degeneration-related defects such as fissures or calcification (Thompson et al., 1990). Discs with a Thompson grade of III or lower were selected for baseline FCD and ion diffusivity measurements. The spines were maintained in fresh-frozen conditions until harvest, and all testing was conducted within 24 hours post-harvest. Motion segments from the C2-C3 (n = 3), C3-C4 (n = 2), C4-C5 (n = 3), C6-C7 (n = 3), and C7-T1 (n = 2, superior site only) vertebral levels were used in the study. IVD segments were cut through the mid-plane using a sterile surgical scalpel, and osteochondral plug specimens containing either NP or AF tissue, along with the underlying layers of CEP and vertebral bone, were extracted. NP, AF, and CEP samples were isolated from these plugs and shaped into 5 mm diameter cylinders, approximately 1 mm thick, for comparative testing (see Figure 1 for details).

Figure 1.

Preparation of IVD tissue specimens. Motion segments from T1 to C2 vertebrae were opened through the mid-plane, and osteochondral plugs containing disc cartilaginous tissue, endplate, and vertebral bone (VB) were isolated. NP and AF disc regions were trimmed to 1.3 ± 0.4 mm and the CEP to 0.9 ± 0.3 mm using a freezing stage microtome (Leica SM2400) prior to conductivity measurements.

Electrical conductivity

Electrical conductivity measurements were employed to evaluate fixed charge density and ion diffusivity within the IVD tissues (Jackson et al., 2009, 2006; Wu et al., 2017; Gu et al., 2002). Freshly prepared samples of NP (n = 7), AF (n = 9), and CEP (n = 7) were immersed in a 0.15 M potassium chloride (KCl) solution and stored at 4°C for 12 hours to equilibrate under isotonic conditions. To prevent swelling and GAG leakage during storage, the tissues were confined peripherally in an acrylic well plate with 5 mm diameter walls and axially between two hydrophilic polyethylene porous platens (50–90 μm pore size, Small Parts, Inc., Miami Lakes, FL) (Figure 2). Electrical conductivity measurements were performed at room temperature (22°C) using a custom apparatus (Figure 2). A constant low-value source current (0.015 mA/cm²) was applied across the IVD samples using a Keithley Sourcemeter (Model 2400, Keithley Instruments, Inc., Cleveland, OH), and axial electrical resistance was measured using a four-wire method. Conductivity was calculated as the ratio of the sample thickness to the product of its electrical resistance and cross-sectional area. Sample thickness was measured during each test using an inbuilt micrometer (Figure 2). This procedure was repeated following a period of equilibration in 0.03 M KCl to assess hypotonic conductivity.

Figure 2.

Schematic of the two-point electrical conductivity method used to measure FCD in cervical IVDs. Co-aligned stainless steel current electrodes and Ag/AgCl voltage sensing electrodes were positioned into a 5 mm nonconductive Plexiglass chamber in contact with samples to measure their electrical resistance. Sample thickness was assessed by a micrometer attached to the upper pair of electrodes, after which electrical conductivity was calculated. Samples were equilibrated in isotonic and hypotonic KCl for 12 hours, prior to each conductivity measurement step.

FCD and Ion Diffusivity

Isotonic and hypotonic electrical conductivity measurements were used to calculate FCD and ion diffusivity following a two-point method (Jackson et al., 2009). Under zero fluid flow conditions, electrical conductivity ($\chi$) is expressed via the Nernst-Einstein relation (Eq. 1) (Helfferich, 1962):

$$\chi =\frac{{F_c}^2{\varphi }^w}{RT}{\sum }_{i=1}^n{z_i}^2{c}_i{D}_i$$ (1)

where $F_c$ is the Faraday constant, ${\varphi }^w$ is the tissue porosity, $R$ is the gas constant, $T$ is absolute temperature, and $c_i$, $z_i$ and $D_i$ are the concentration, valence, and diffusivity of mobile ions. For ${\mathrm{K}}^{+}(\mathrm{z}=+1,c^{+})$ and ${\text{Cl}}^{\text{-}}(\mathrm{z}=\text{-},c^{-})$ ions from a KCl bath (Jackson et al., 2009), Eq. 1 simplifies to:

$$\chi =\frac{{F_c}^2{\varphi }^w}{RT}(c^{+}{D}^{+}+c^{-}{D}^{-})$$ (2)

Assuming electroneutrality $(c^{+}=c^{-}+c^F)$ (Lai et al., 1991), where $c^F$ denotes the FCD of the tissue, and Donnan equilibrium conditions $(c^{+}{c}^{-}={(c^{\ast })}^2)$ (Maroudas, 1968, 1975), where $c^{\ast }$ denotes the KCl bath concentration, the mobile ion concentrations can be solved and substituted into Eq. 2, yielding:

$$\chi =\frac{{F_c}^2{\varphi }^w}{RT}D\sqrt{{(C^F)}^2+4{(C^{\ast })}^2}$$ (3)

Here, $D$ is the ion diffusivity, assumed equal for K⁺ and Cl⁻, based on their similar Stokes radii (K⁺: r_s = 0.137nm; Cl⁻: r_s = 0.142nm) (Gu et al., 2004). Since the tissue samples are confined and cannot swell under varying KCl concentrations, FCD and tissue porosity remain constant. From Eq. 3, $c^F$ may be isolated and equated under the different KCl bath conditions tested, enabling direct calculation of FCD from experimental parameters (Eq. 4):

$$c^F=2\sqrt{\frac{{\chi }_{iso}^2[{(c_{iso}^{\ast })}^2-{(c_{hypo}^{\ast })}^2]}{{\chi }_{iso}^2-{\chi }_{hypo}^2}-{(c_{iso}^{\ast })}^2}$$ (4)

where ${\chi }_{iso}$ and ${\chi }_{hypo}$ represent the measured isotonic and hypotonic conductivities, and $c_{iso}^{\ast }$ and $c_{hypo}^{\ast }$ denote the equilibrated KCl concentrations. Ion diffusivity can also be obtained through manipulation of Eq. 3 assuming constant FCD (Eq. 5):

$$D=\frac{RT}{2F_c^2{\varphi }^w}\sqrt{\frac{{\chi }_{iso}^2-{\chi }_{hypo}^2}{{(c_{iso}^{\ast })}^2-{(c_{hypo}^{\ast })}^2}}$$ (5)

Biochemical Assays

Porosity was determined using a buoyancy-weighing method (Gu et al., 1996, 2002; Yao et al., 2002), with the aid of an analytical balance (precision of 100 μg) and a density-determination kit (Sartorius, Germany). Porosity was calculated from specimen weights measured initially in air (${\mathrm{W}}_{\text{wet}}$), submerged in 1x PBS saline solution (${\mathrm{W}}_{\text{PBS}}$) after conductivity experiments, and in air again after lyophilization for one week (${\mathrm{W}}_{\text{dry}}$). The relationship between these measurements is given by Eq. 6:

$${\varphi }^w=\frac{W_{wet}-W_{dry}}{W_{wet}-W_{PBS}}\ast \frac{{\rho }_{PBS}}{{\rho }_w}$$ (6)

Where the ratio $\frac{{\rho }_{PBS}}{{\rho }_w}=1.005$ represents that of the specific weights of the PBS solution and water. After lyophilization, glycosaminoglycan (GAG) content in NP, AF, and CEP tissues was quantified using the Blyscan^™ Glycosaminoglycan assay kit (Biocolor Life Science, Northern Ireland). This assay measures GAG content through the absorbance of 1,9-dimethyl-methylene blue dye bound to GAG molecules. Additionally, small tissue portions unsuitable for conductivity analysis due to their shape were included in the regional dry weight GAG characterization (NP: n = 18; AF: n = 21; CEP: n = 15 assays total).

Histological Characterization

Two cervical IVD segments (C2-C3 and C3-C4 levels) were both sectioned in the sagittal and coronal planes (~4 mm thick). Tissue was fixed overnight in 10% neutral buffered formalin (Sigma-Aldrich), decalcified using a 10% ethylenediaminetetraacetic acid (EDTA) solution (Sigma-Aldrich), and subsequently embedded in paraffin wax. Serial tissue sections (7 μm) were prepared on slides and stained with Safranin-O and Fast Green, Ehrlich’s hematoxylin and eosin (H&E), and Picrosirius Red. Safranin-O staining was used to visualize proteoglycan content, while H&E staining provided detailed examination of IVD tissue morphology. Bright-field images were captured using a 10x objective on a BZ-X810 microscope (Keyence, Itasca, IL). Picrosirius Red-stained sections were imaged under polarized light using a 20x objective on an Olympus BX53 microscope (Olympus, Center Valley, PA) to highlight birefringent collagen fibers. Quantitative analysis of Safranin-O stain intensity by color deconvolution of the images, as well as measurement of fiber orientation from the Picrosirius red images was performed using Fiji ImageJ software (Schindelin et al., 2012) (see supplement for details).

Statistical Analysis

Electrical conductivity (under isotonic and hypotonic conditions), fixed charge density, and ion diffusivity were compared across the different disc regions (NP, AF, and CEP). Linear mixed effects models were fit to each parameter using R software (Pinheiro et al., 2024; R Core Team, 2024), with spine donor included as a random effect. Due to sample size constraints, the effects of aging were not considered. Regional tissue properties were estimated using a restricted maximum likelihood approach and summarized as model-based averages with 95% confidence intervals (mean ± SD reported in Table 1). Type III ANOVA tests were conducted to evaluate overall regional variability in the IVD properties, while pairwise comparisons between NP, AF, and CEP were performed using a generalized t-test procedure with Bonferroni-Holm p-value correction (Hothorn et al., 2008; Holm, 1979). Pearson correlations were also calculated to assess the relationship between FCD and GAG content, as well as between ion diffusivity and porosity. A significance threshold of p < 0.05 was used for all statistical tests.

Table 1.

Fixed charge density and ion diffusivity properties of human cervical and lumbar IVDs, bovine tail IVDs, and human articular cartilage (AC). *Reported based on ratio of water and tissue weights, not porosity. **Wet weight GAG content estimated by combination of reported dry weight and water content results. ***Indicates an average of values from different degeneration stages (scores I-III considered as relevant baseline parameters).

Tissue	FCD (mEq/g wet tissue)	Ion Diffusivity (10−6 cm2/s)	Porosity (${\varphi }^{\mathit{W}}$)	GAG (μg/mg wet tissue)	GAG (μg/mg dry tissue)	Reference
Cervical CEP	0.26 ± 0.14	6.34 ± 1.70	0.63 ± 0.17	60.6 ± 12.0	126.4 ± 59.8	This study
Cervical NP	0.17 ± 0.07	10.71 ± 2.54	0.86 ± 0.04	37.2 ± 6.8	191.8 ± 87.0	This study
Cervical AF	0.13 ± 0.03	11.88 ± 2.39	0.85 ± 0.06	32.1 ± 8.3	183.0 ± 70.1	This study
Cervical CEP	-	-	-	-	~ 231***	(Tomaszewski et al., 2015)
Cervical NP	-	-	-	-	~ 795***	(Tomaszewski et al., 2015)
Cervical AF	-	-	-	-	~ 615***	(Tomaszewski et al., 2015)
Cervical NP	-	-	-	-	169.9 ± 37.3	(Bostelmann et al., 2017)
Cervical AF	-	-	-	-	132.4 ± 42.2	(Bostelmann et al., 2017)
Cervical NP	-	-	~ 0.77*	~ 57.5**	~ 250	(Scott et al., 1994)
Cervical AF	-	-	~ 0.71*	~ 69.6**	~ 240	(Scott et al., 1994)
Lumbar CEP	0.12 ± 0.03	2.97 ± 1.00	0.73 ± 0.07	31.2 ± 5.1	93.6 ± 18.4	(Wu et al., 2017)
Lumbar NP	0.18 – 0.28	-	~ 0.82	-	-	(Urban and Maroudas, 1979)
Lumbar AF	0.07 – 0.13	-	~ 0.60	-	-	(Urban and Maroudas, 1979)
Bovine NP	0.19 ± 0.04	10.3 ± 1.6	0.93 ± 0.02	~34.3**	490 ± 64	(Jackson et al., 2009)
Bovine AF	0.06 ± 0.03	7.56 ± 2.12	0.81 ± 0.05	~10.3**	54 ± 41	(Jackson et al., 2009)
Human AC	0.10 – 0.25	6.9 – 7.8	-	-	-	(Maroudas, 1968)
Human AC	0.02 – 0.15	-	-	-	-	(Hasegawa et al., 1983)

Results

Electrical Conductivity

Electrical conductivity under isotonic conditions varied significantly by IVD region (p = 0.0003), whereas this effect was not statistically apparent under hypotonic conditions (p = 0.0619). In both conditions, the conductivity of the CEP (isotonic: 7.7 mS/cm, 95% CI [5.8, 9.5]; hypotonic: 5.7 mS/cm, [3.7, 7.7]) was significantly lower than that of the NP (isotonic: 12.6 mS/cm, [10.7, 14.4], p < 0.0001; hypotonic: 7.7 mS/cm, [5.8, 9.7], p = 0.0321) (Figure 3). Additionally, under isotonic conditions, CEP conductivity was lower compared to AF (13.0 mS/cm, [11.3, 14.6], p < 0.0001) (Figure 3). Conductivity did not differ significantly between the NP and AF regions under either condition.

Figure 3.

Regional baseline electrical conductivities of human cervical IVDs under isotonic and hypotonic conditions. Asterisks indicate differences at the given levels of statistical significance (* p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001).

FCD and Ion diffusivity

Both FCD and ion diffusivity varied significantly by tissue region (p = 0.0203 and p = 0.0002, respectively). FCD was highest in the CEP (0.24 mEq/g wet tissue, [0.16, 0.33]) and was significantly greater than in the AF (0.14 mEq/g wet tissue, [0.07, 0.22], p = 0.0060), though no significant difference was observed between the CEP and NP (Figure 4). Ion diffusivity exhibited an inverse trend, being significantly lower in the CEP (6.3 × 10⁻⁶ cm²/s, [4.5, 8.1]) compared to the NP (10.7 × 10⁻⁶ cm²/s, [8.9, 12.5], p = 0.0002) or AF (11.9 × 10⁻⁶ cm²/s, [10.3, 13.5], p < 0.0001) (Figure 4). No significant differences in FCD or ion diffusivity were found between the NP and AF regions.

Figure 4.

(a) Fixed charge density and (b) ion diffusivity quantified in human cervical IVDs. Asterisks indicate differences at the given levels of statistical significance (* p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001).

GAG and Porosity

Significant regional variation was observed in GAG content (p = 0.0003) and porosity (p = 0.0003). The CEP exhibited the highest GAG content (58.8 μg/mg wet tissue, [46.7, 70.9]), which was significantly greater than that of the NP (37.8 μg/mg wet tissue, [27.7, 48.0], p < 0.0001) or AF (34.0 μg/mg wet tissue, [24.7, 43.3], p < 0.0001). The CEP also had the lowest porosity (0.63 [0.55, 0.71]), significantly lower than that of both the NP (0.86 [0.78, 0.94], p < 0.0001) and AF (0.85 [0.78, 0.93], p < 0.0001). No significant differences were found in GAG content or porosity between the NP and AF.

Correlation of FCD and Ion diffusivity with biochemical composition

A strong positive correlation was observed between FCD and GAG content (r = 0.65, p = 0.0047) (Figure 5). Additionally, ion diffusivity showed a moderate but significant positive correlation with porosity (r = 0.53, p = 0.0097) (Figure 5).

Figure 5.

(a) Correlation of fixed charge density with glycosaminoglycan content and (b) correlation of ion diffusivity with porosity.

Histological Appearance

Histological sections of the cervical IVD stained with Safranin-O and Fast Green, as well as H&E, revealed distinct layers of vertebral bone, disc, and CEP tissue, with the CEP measuring approximately 0.9 mm in thickness (Figure 6a & b). In Safranin-O-stained sections, the CEP displayed a deep red hue, characteristic of higher proteoglycan content compared to the NP and AF regions. This observation was further supported by color deconvolution analysis of the Safranin-O stain, where lower gray pixel values in the CEP confirmed a stronger staining intensity (Figure 6d and Supp. Figure 1). Picrosirius Red-stained sections viewed under polarized light exhibited a compact, transversely oriented collagen layer in the CEP (Figure 6c & e, and Supp. Figure 1). By contrast, collagen fibers in the AF were organized with a slightly off-axial orientation (Figure 6c & e, and Supp. Figure 1). The NP, while more disorganized and lacking a preferred fiber orientation, displayed a fibrous structure (Figure 6c & e).

Figure 6.

Representative histology of cervical IVD tissue regions, including (a) Safranin-O and Fast Green, (b) Hematoxylin and Eosin, and (c) Picrosirius Red stains from the C2-C3 level. Additionally, regional quantification of (d) Safranin-O stain intensity, based on color deconvolution of images in (a), and (e) collagen fiber orientation, analyzed from grayscale images in (c), regions of interest measured 442 × 460 μm.

Discussion

Fixed charge density quantified in cervical discs provides important comparisons to data from prior studies on lumbar IVDs, animal models, and other human cartilaginous tissues (Table 1). Notably, the FCD in the cervical NP was lower than in the lumbar NP (Urban and Maroudas, 1979). As FCD contributes directly to the extracellular matrix osmotic pressure, this result suggests that the cervical nucleus may experience less swelling than its lumbar counterpart (Lai et al., 1991; Urban and McMullin, 1988; Urban and Maroudas, 1979). Consequently, a lower FCD in the cervical NP may also be associated with reduced matrix stiffness in compression (Cortes et al., 2014; Urban and McMullin, 1988), although further biomechanical testing is required to confirm this. The reduced FCD in the cervical NP may reflect lower weight-bearing requirements of cervical discs overall, which support only the weight of the head and share compressive loads with zygapophysial joints positioned laterally along the vertebral arch (Bogduk and Mercer, 2000; Busscher et al., 2010; Kumar and Pai, 2020; Galbusera and Wilke, 2018). Mechanically, intact cervical disc segments exhibit an ultimate compressive strength that averages approximately 45% of that in lumbar discs (Przybyla et al., 2007).

The relatively more homogeneous FCD distribution between the NP and AF in cervical discs is also intriguing (Table 1 and Supp. Table 1), and contrasts with lumbar discs, where FCD in the NP is 2-3 times higher than in the AF (Urban and Maroudas, 1979). A reduced osmotic gradient between the cervical NP and AF, resulting from more uniform FCD distribution, could reduce internal resistance to bending or torsion (Przybyla et al., 2007). Consistent with this observation, radiographic studies of cervical discs have demonstrated greater ranges of flexion and axial rotation compared to lumbar segments (Bogduk and Mercer, 2000; Nakanishi et al., 2024). Cervical discs also feature a unique saddle-shaped geometry and a thinner crescentic-shaped AF that does not fully encapsulate the NP posteriorly, considered to be more optimal for spinal mobility (Mercer and Bogduk, 1999; Bogduk and Mercer, 2000; Kim et al., 2017; Galbusera and Wilke, 2018). Regarding the thinner-walled features of the cervical AF, relative homogeneity in the osmotic environment may help to prevent herniations through reduced tensile stress concentration at the NP-AF interface (Chahine et al., 2004; Ateshian et al., 2009). A comparatively lower fluid pressure in the cervical NP driven by lower FCD could further avoid endplate injuries like Schmorl’s nodes which are common in lumbar discs, and understood to disrupt nutrient availability (Adams and Hutton, 1982; Soukane et al., 2007). Moreover, lower NP fluid pressure could result in reduced fluxes of nutrients overall with loading. In addition to FCD, this study also quantified ion diffusivity for potassium (K⁺) and chloride (Cl⁻) ions in the cervical IVD, revealing regional variations. As with FCD, ion diffusivity did not differ significantly between the NP and AF. The relative uniformity in ion diffusivity in the cervical disc could be explained by the more fibrocartilaginous morphology of the cervical nucleus, which was revealed by Picrosirius Red histology imaging (Figure 6c), in agreement with literature (Tonetti et al., 2005; Scott et al., 1994; Oda et al., 1988).

FCD in the cervical CEP measured nearly double the value reported for lumbar CEP tissue (Table 1). In cervical discs, which experience greater mobility and likely higher frequency of loading, this heightened FCD may further limit fluid convection across the IVD-vertebral bone interface, helping to stabilize nutrient gradients within the disc (Ferguson et al., 2004; Yao and Gu, 2004; Zhu et al., 2012). The higher FCD in the CEP may also provide mechanical protection by mitigating endplate fracture risk in high-strain environments (Adams and Hutton, 1982), however, literature on cervical endplate fracture patterns specifically is sparse. Compared to other disc regions, the elevated FCD in the CEP highlights its significance as a mechanical barrier, supporting fluid pressurization within the disc at any spinal level (Buchweitz et al., 2024; Wu et al., 2017). The lower ion diffusivity observed in the CEP compared with NP and AF, also suggests that this region is more restrictive to solute transport. This finding aligns with lumbar disc studies, where lower porosity and solute diffusivities in the CEP are thought to protect the IVD from rapid solute fluxes during compression, even under dynamic loading (Wu et al., 2017; Yao and Gu, 2004; Wu et al., 2013). Interestingly, ion diffusivity in the cervical CEP was greater than previously reported for the lumbar CEP (Table1). Combined with the shorter diffusion distances resulting from smaller disc heights, increased solute diffusivity in the CEP could aid nutritional recovery in cervical discs, particularly in the context of a dynamic loading environment where nutrient gradients are more affected by fluid convection (Yao and Gu, 2004).

The positive correlation observed between FCD and GAG content in the cervical IVD is consistent with lumbar IVD studies, confirming that proteoglycan molecules remain primary determinants of FCD in cervical disc tissues (Jackson et al., 2009; Wu et al., 2017). However, a slight discrepancy was noted between regional FCD and GAG distributions: while both properties follow similar trends, FCD in the CEP was significantly greater than in the AF but not the NP, whereas GAG content was significantly greater in the CEP relative to both regions (Wu et al., 2017). This could be due to regional variations in GAG types, such as keratan sulfate and chondroitin sulfate, which carry different charges per disaccharide, and could have differing influence on the regional FCD pattern (Jackson et al., 2009; Roberts et al., 1994; Stevens et al., 1979; Wu et al., 2017). Safranin-O histology images align with both the regional GAG distribution and FCD quantification, as the CEP exhibited a stronger red hue compared to other disc regions, indicating greater proteoglycan content (Figure 6a & d, Supp. Figure 1). The correlation between ion diffusivity and porosity also indicates that cervical IVD water content significantly influences ion transport, although the strength of this relationship may vary by region, particularly in the CEP, where tissue was least porous, and the association was weaker. Correlation of ion diffusivity with porosity was expected, as the calculation of ion diffusivity is at least in part dependent on porosity (Eq. 5).

The findings of this study may have important applications for both cervical IVD tissue engineering and allograft preservation. Tissue-engineered discs are designed to replace damaged IVD tissue, providing mechanical stability and flexibility outcomes superior to those of fusion surgeries (Hudson et al., 2013; Bowles et al., 2012). Given the critical role of FCD in the compressive response and osmotic environment of the IVD (Cortes et al., 2014; Gu and Yao, 2003), the regional characterization of FCD from this study provides essential electro-mechanical benchmarks for developing physiologically relevant scaffolds (Nerurkar et al., 2011; Thomas et al., 2024). These scaffolds should incorporate both FCD and proteoglycan content to better mimic cervical disc biomechanics, which support higher mobility demands. For allograft preservation, maintaining elevated FCD in the CEP may be key to preserving electro-mechanical properties at the IVD-bone interface, thereby improving graft longevity and functionality (Shalash et al., 2024; Chan et al., 2010; McCarty et al., 2010; Ruan et al., 2007). Additionally, the findings of this study support the advancement of computational models for cervical IVD mechanics and transport dynamics, which are critical for understanding disc degeneration and pathophysiology (Zhou et al., 2021; Jackson et al., 2011; Wu et al., 2013).

While this study provides insights into the electro-biomechanical properties of cervical IVDs, several limitations must be acknowledged. The sample size was small (n = 7–9 per region from N = 5 human subjects), limiting the generalizability of FCD results. Additionally, with the inclusion of grade III discs, early degeneration was present among this cohort. FCD and ion diffusivity were quantified solely as baseline measurements, and did not account for the effects of sex, age, disc level, or degeneration progression (Urban and McMullin, 1988). While dry-weight GAG measurements in this study align with previously reported values for cervical disc tissues, they are lower than those reported by Tomaszewski et al. (2015) (Table 1). This discrepancy is possibly attributed to the use of frozen rather than freshly excised tissue (within 24 hours post-mortem). Regionally, however, the ratio of GAG content between NP and AF was in fair agreement across prior cervical IVD studies (Supp. Table 1). Variations in sample thickness, particularly between the CEP and other disc components (approximately 400 μm), could introduce variability in conductivity measurements, affecting FCD outcomes. Future studies with larger and more diverse populations will be essential to validate these findings and explore how cervical disc FCD and ion diffusivity change with aging and degeneration.

This study provides the first regional characterization of FCD in cervical IVDs. Given the close relationship between FCD and osmotic pressure, lower FCD in cervical NP compared to lumbar NP tissue aligns with the reduced weight-bearing capacity of cervical discs. A more homogeneous FCD distribution between cervical NP and AF (Table 1 and Supp. Table 1) is also unique and suggests greater uniformity in the cervical disc osmotic environment, beneficial to bending and torsional compliance. Elevated FCD in the cervical CEP compared to lumbar CEP, plays a potential role in endplate injury prevention, particularly with increased cervical disc mobility. Greater FCD in the CEP relative to the NP and AF also confirms the importance of the CEP as a mechanical barrier to the IVD at any spinal level (Wu et al., 2017). These findings advance our current understanding of the cervical disc electro-mechanical environment and may offer essential benchmarks for cervical disc biomechanical modeling and other tissue-level characterizations.