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Anatomical and Micro-CT measurement analysis of ocular volume and intraocular volume in adult Bama Miniature pigs, New Zealand rabbits, and Sprague-Dawley rats
Different species of animal eye volume
Wu Yajun Writing – original draft Writing – review & editing 1 2 3
Feng Yuliang Writing – review & editing 1 2 3
Yang Jiasong Data curation Investigation 1 2 3
Ran Yuwen Investigation Methodology Visualization 4
Shu Zongtao Formal analysis Methodology 5
Cen Xiaobo Project administration Resources 5 *
https://orcid.org/0000-0002-1172-3349
Li Wensheng Project administration Supervision 1 2 3 *
1 Aier Academy of Ophthalmology, Central South University, Changsha, Hunan, China
2 Department of Ophthalmology, Shanghai Aier eye hospital, Shanghai, China
3 Shanghai Aier eye institute, Shanghai, China
4 Changsha Aier eye hospital, Changsha, Hunan, China
5 WestChina-Frontier PharmaTech Co.,Ltd., Chengdu, Sichuan, China
Goździewska-Harłajczuk Karolina Editor
Wrocław University of Environmental and Life Sciences: Uniwersytet Przyrodniczy we Wroclawiu, POLAND
Competing Interests: NO authors have competing interests.

* E-mail: drlws@qq.com (WL); xbcen@scu.edu.cn (XC)
20 9 2024
2024
19 9 e031083017 7 2024
6 9 2024
© 2024 Wu et al
2024
Wu et al
https://creativecommons.org/licenses/by/4.0/ This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Aim

Utilizing a combination of micro-computed tomography (micro-CT) and anatomical techniques for the volumetric assessment of the eyeball and its constituents in Bama Miniature Pigs, New Zealand rabbits, and Sprague-Dawley(SD) rats.

Method

Six Bama Miniature pigs, New Zealand rabbits, and SD rats were enrolled in the study. Micro-CT and gross volumetric estimation of ocular volume were employed to acquire data on ocular volume, anterior chamber volume, lens volume, and vitreous cavity volume for each eye.

Results

The eyeball volume of pigs ranges from approximately 5.36 ± 0.27 to 5.55 ± 0.28 ml, the lens volume from approximately 0.33 ± 0.02 to 0.37 ± 0.06 ml, the anterior chamber volume from approximately 0.19 ± 0.05 to 0.28 ± 0.04 ml, and the vitreous volume is approximately 3.20 ± 0.18 ml. For rabbits, the eye volume, lens volume, anterior chamber volume, and vitreous volume range from approximately 3.02 ± 0.24 to 3.04 ± 0.24 ml, 0.41 ± 0.02 to 0.44 ± 0.02 ml, 0.23 ± 0.04 to 0.26 ± 0.05 ml, and 1.54 ± 0.14 ml, respectively. In SD rats, the volumes are 0.14 ± 0.02 to 0.15 ± 0.01 ml for the eyeball, 0.03 ± 0.00 to 0.03 ± 0.00 ml for the lens, 0.01 ± 0.00 to 0.01 ± 0.01 ml for the anterior chamber, and 0.04 ± 0.01 ml for the vitreous volume.

Conclusion

The integration of micro-CT and gross volumetric estimation of ocular volume proves effective in determining the eyeball volume in Bama Miniature Pigs, New Zealand rabbits, and SD rats. Understanding the volume distinctions within the eyeballs and their components among these experimental animals can lay the groundwork for ophthalmology-related drug research.

Science Research Foundation of Aier Eye Hospital Group AR2201D3 2023 Research Fund of Aier Eye Research Institute AEI202310LC01 https://orcid.org/0000-0002-1172-3349
Li Wensheng 2023 Research Fund of Aier Eye Research Institute(No.AEI202310LC01); Science Research Foundation of Aier Eye Hospital Group (No.AR2201D3). 1.2023 Research Fund of Aier Eye Research Institute (No.AEI202310LC01), Funder: Wensheng Li:Grant project funding, guide research design, and revise manuscripts. 2.Science Research Foundation of Aier Eye Hospital Group (No.AR2201D3), Funder: Jiasong Yang: Grant project funding, responsible for data checking and statistics. Data AvailabilityAll relevant data are within the manuscript and its Supporting information files.
Data Availability

All relevant data are within the manuscript and its Supporting information files.
==== Body
pmcIntroduction

The eyeball, a pivotal sensory organ with a precise refractive system, plays a crucial role in the human body [1]. Biomechanical processes involving fluid flow among different ocular structures are vital for delivering ophthalmic drugs [2]. Thus, elucidating the eyeball and its contents’ volumes holds fundamental scientific value. This encompasses research on emmetropia and refractive errors mechanisms, establishing eye biological models, and contributes significantly to the diagnosis and treatment of various eye diseases.

Pigs, with eyes structurally and sizably similar to humans, are frequently employed as animal models for eye disease research [3–5]. Notably, studies using pigs, such as Chan et al. [6] have used pig eyes to validate the effectiveness of gene therapy. In addition, New Zealand rabbits, known for their docile nature and large eyes, are also prevalent in ophthalmic research [7, 8]. Sun et al. [9] utilized them for microwave thermokeratoplasty experiments. Besides, Rabbits, characterized by larger eyeballs and lacrimal glands akin to humans, are widely utilized in dry eye studies due to their stability [10]. Moreover, rabbits have large eyes that are conducive to the biochemical characteristics of drug research, and their usage cost is relatively low, though non-human primates are more similar to humans, their usage cost is very high, making rabbits a more commonly used choice for dry eye experiments [11]. Additionally, Sprague-Dawley(SD) rats, cost-effective with rapid reproduction, share a comparable eyeball structure with humans. Guo, et al., etc [12]. successfully established an animal model of diabetes by intraperitoneal injection of streptozotocin (STZ) into SD rats, and observed abnormal meibomian gland function in model rats, revealing that diabetes can cause dysfunction of the meibomian gland [12]. In conclusion, experimental animals, including Bama miniature pigs, New Zealand rabbits, and SD rats, are pivotal in advancing our understanding of human eye diseases in ophthalmology research.

Computed tomography (CT) imaging provides detailed visualization of the eyeball, offering high resolution and measurement accuracy. Qualitative and quantitative assessments of specific data can be conducted through three-dimensional reconstruction [13, 14]. Micro-CT, a miniature CT with micrometer-level resolution, enhances spatial resolution for imaging small animal disease models [15]. While Magnetic Resonance Imaging (MRI) excels in finer soft tissue imaging, its higher cost and longer scanning time, coupled with limitations in scanning metallic or paramagnetic tissues [16], especially in animal experiments, few laboratories have the conditions to equip Micro-MRI systems. Moreover, research has consistently demonstrated CT’s efficacy in measuring ocular volume parameters. In fact, as early as 1984, one study used CT to measure the ocular volume of the human body to study its correlation with age and gender [17]. Besides, Ozer CM, et al. [18] also clearly indicated the important value of CT in measuring eyeball volume parameters. Also, CT is used in animal experiments, Leszczyński B, et al. [14] performed CT scans on the eyeballs of domestic pigs and combined staining methods to accurately obtain structurally clear pig eye tissue. Similarly, Salguero et al. [19] measured the volume, density, and the length of the normal dog eye structures (Including the average axial length of the sphere, the average anterior posterior distance of the anterior chamber, the average anterior posterior distance of the vitreous chamber, etc) with CT.

Despite these advancements, uncertainties persist regarding specific parameters of animal eyeballs and whether imaging data align with actual values. In this study, we utilized micro-CT combined with anatomical methods to simultaneously measure ocular volume and intraocular volume in three common ophthalmic experimental species-Bama miniature pigs, New Zealand rabbits, and SD rats. This approach aims to obtain more accurate parameters for the eyeball and intraocular volume in these species, providing crucial reference values for related ophthalmic experiments.

Method

Ethics

All animals in this study were approved by the Institutional Animal Care and Use Committee (IACUC) of WestChina-Frontier Pharma Tech Co., Ltd. (Approval Number: IACUC- SW-S2023031-P001-01). Our experiment comply with the following animal research guidelines: Animal Research: Reporting in Vivo Experiments (ARRIVE) guidelinesthe, Association for Research in Vision and Ophthalmology (ARVO), and the American Veterinary Medical Association (AVMA) Guidelines for the Europe of Animals (2020).

Experimental animals

We selected 6 healthy Bama miniature pigs (ordinary grade), aged 10–12 months, weighing 28–33 kg, with an equal gender distribution. The pigs were obtained from Chengdu Dashuo Experimental Animal Co., Ltd. (License: SCXK [Chuan] 2019–031). Additionally, 6 SPF grade SD rats (half male, half female), aged 2–3 months, weighing 330-420g, were purchased from Zhejiang Weitong Lihua Experimental Animal Technology Co., Ltd. (License: SCXK [Zhe] 2019–0001). The cohort also included 6 SPF grade New Zealand rabbits (half male, half female), 7 months old, weighing 3.0–3.5kg, sourced from Shandong Benming Biotechnology Co., Ltd. (License: SCXK [Lu] 2017–003). Each species of animal includes a total of 12 eyeballs. All animals were housed at WestChina-Frontier Pharma Tech Co., Ltd. (License for experimental animal use: SYXK [Chuan] 2021–238). The animals were maintained in a controlled environment with a temperature range of 16–26°C for rabbits and pigs and 20–26°C for rats (daily temperature difference ≤ 4°C), relative humidity of 40–70%, and a 12/12-hour light/dark cycle.

Micro-CT scanning and data analysis

We utilized the NEMO® Micro-CT (model NMC-100 by Life Medical Technology, Kunshan, China) for scanning. Scanning parameters were set as follows: large detached cabin for scanning; current of 0.2mA; voltage of 70kV; 40 frames per second; 360 frames per turn; and a scanned pixel size of 0.03*0.03*0.03mm. Reconstruction parameters were configured with an iterative algorithm, a reconstruction separation rate of 1k*1k, and a reconstruction pixel size of 0.015*0.015*0.015mm.

Volumetric measurements and image analysis:We employed Avatar software (version 1.6.6.7, provided by Pingsheng Medical Technology, Kunshan, China) for image analysis. The CT reconstructed image was imported into Avatar. Using the [Hand-drawn Mask] tool, the specific way we locate the eyeball for scanning is to ensure that the entire eyeball is located within the scanning interval, manually select the lens to start drawing from the tip of the ciliary body, move towards the other end until the lens structure disappears, and then use this tool to draw the region of interest (ROI) of the lens until the anatomical structure disappears; Using the same method, select any starting surface of the eyeball or anterior chamber structure and draw its ROI until the anatomical structure disappears, the vitreous body and posterior chamber cannot be located due to the lack of a clear boundary line. Each ROI can be divided into multiple sections when drawing, and the software automatically calculates the ROI volume of the middle section. After the ROI of each eye structure such as anterior chamber and lens was drawn, Avatar software provides mask of the lens and anterior chamber, and automatically calculates its volume as the physiological volume of the corresponding structure (see Fig 1), without secondary or manual calculation.

10.1371/journal.pone.0310830.g001 Fig 1 CT images of the eyeballs and contents of different species of animals after manual delineation.

Note: A, CT images of the eyeballs and contents of adult Bama miniature pigs; B, CT images of the eyeballs and contents of adult New Zealand rabbits; C, CT images of the eyeballs and contents of adult SD rats.

The total eyeball volume was determined using the threshold selection method. The vitreous body volume was calculated using Boolean operations based on the difference between the total eyeball volume and the lens and anterior chamber volume. It is important to note that the volume of the vitreous cavity obtained here represents the sum of the vitreous cavity and the remaining eye tissue, as the CT images lack clarity in delineating the vitreous cavity and posterior chamber. The actual volume of the vitreous cavity was measured using anatomical methods.

All operations were performed by a single experimenter (ZTS).

Gross volumetric estimation of ocular volume methods

Pigs and rabbits: After intramuscular injection of 2mg/kg of diazepam (Sigma-Aldrich, USA), intravenous injection (via ear vein after skin preparation) of pentobarbital sodium (30mg/kg, Sigma-Aldrich, USA) was administered, and ultimately all animals were euthanized through abdominal aortic bleeding. Only when the animal’s breathing, heartbeat, and toe reflexes completely disappear can it be confirmed that the animal has died, and rapid enucleation of the eyeball can be performed. Both eyeballs were removed and placed in a 10ml measuring cup. Subsequently, 3ml of distilled water was added incrementally using 0.5ml, 0.3ml, and 0.1ml pipettes until reaching the 10ml mark. The total transferred water amount was recorded to determine the total ocular volume. A 1ml syringe was used to extract aqueous humor until the anterior chamber was dry, and the volume of the aqueous humor was recorded. After aqueous humor removal, the eyeball underwent rapid freezing in liquid nitrogen, and the complete lens and vitreous body were dissected. These tissues were then cut into pieces and placed into a 5ml measuring cylinder. Following tissue melting, 0.5ml, 0.3ml, and 0.1ml pipettes were used to inject distilled water into the measuring cup until reaching the 5ml mark. The total transferred water amount was recorded to obtain the volume of the vitreous body and lens.

Rats: The measurement method for the eyeball volume of SD rats is the same as that of pigs and rabbits. Unlike euthanasia and dissection of pigs and rabbits using intravenous anesthesia, SD rats were anesthetized with 30mg/kg pentobarbital sodium (Sigma-Aldrich, USA) via intraperitoneal injection, followed by bloodletting of the abdominal aorta and enucleation of the eyeball. Besides, unlike the 10ml measuring cup used for the other two types of eyeballs, the eyeballs of rats were placed in a 5ml measuring cup to determine ocular volume. A 0.3ml insulin syringe was used for extracting aqueous humor. The lens and vitreous body removal method remained the same as in rabbits and pigs. Extracted crystals and vitreous bodies were placed into the 0.1ml scale of the syringe for insulin, and the volume was read after tissue melting.

All the aforementioned procedures were conducted by a single experimenter.

Statistical methods

Statistical analysis of the data was performed using SPSS 25.0 (IBM Corp., Armonk, NY, USA). All results are presented as mean±standard deviation (SD). A comparison of volumes measured by the two methods was carried out using unpaired t-tests, Pearson correlation analysis was performed using the software Origin (version 2022, Originlab, USA), and p≤0.05 indicated a statistically significant difference.

Result

Results of ocular volume and intraocular contents in Bama Miniature Pigs

No significant difference was observed between the anatomical measurement data and CT data of ocular volume and vitreous cavity volume in Bama miniature pigs (p > 0.05), as illustrated in Fig 2A and 2D. However, the gross volumetric estimation of ocular volume indicated greater lens volume and anterior chamber volume compared to CT results, as depicted in Fig 2B and 2C, respectively. In addition, the eyeball volume of pigs was about 5.36±0.27 to 5.55±0.28ml, the lens volume was about 0.33±0.02 to 0.37±0.06ml, and the anterior chamber volume was about 0.19±0.05 to 0.28±0.04ml. The vitreous volume was about 3.20±0.18ml. Refer to Table 1 for detailed data.

10.1371/journal.pone.0310830.g002 Fig 2 Comparison of anatomical and CT methods for measuring the eyeball and intraocular volume in Bama miniature pigs (ml).

Note: A, Comparison of two measurement methods in pig ocular volume; B, Comparison of two measurement methods in pig lens volume; C, Comparison of two measurement methods on the volume of pig anterior chamber; D, Comparison of two measurement methods on the volume of pig vitreous cavity (The volume of the vitreous cavity here represents the total volume of the vitreous cavity and the remaining eye tissue). ns means that the difference is not statistically significant; *Indicates P ≤ 0.05; *** Indicates P ≤ 0.001.

10.1371/journal.pone.0310830.t001 Table 1 Comparison of anatomical and CT measurements of ocular volumes and intraocular contents volumes in pigs (ml).

Group (n = 12)	Ocular volume	Lens volume	Anterior chamber volume	Vitreous cavity volume (volume of vitreous cavity and remaining eye tissue)	Real vitreous cavity volume	
Gross volumetric estimation of ocular volume	5.55±0.28	0.37±0.06	0.28±0.04	4.91±0.29	3.20±0.18	
CT result	5.36±0.27	0.33±0.02	0.19±0.05	4.86±0.23	/	
t value	1.66	2.15	4.45	0.40	/	
P value	0.11	0.04*	≤ 0.00***	0.69	/	
Note:

*Indicates P ≤ 0.05;

*** Indicates P ≤ 0.001.

Results of ocular volume and intraocular contents in New Zealand rabbits

Similar to the findings in Bama miniature pigs, there was no significant difference between the gross volumetric estimation of ocular volume and CT results for ocular volume and vitreous cavity volume in New Zealand rabbits (p>0.05), as depicted in Fig 3A and 3D, respectively. However, the gross volumetric estimation of ocular volume of lens volume and anterior chamber volume were notably higher than the CT results, as illustrated in Fig 3B and 3C, respectively. The eyeball volume of rabbits was about 3.02±0.24 to 3.04±0.24ml, the lens volume was about 0.41±0.02 to 0.44±0.02ml, the anterior chamber volume was about 0.23±0.04 to 0.26±0.05ml, and the vitreous volume was about 1.54±0.14ml. Detailed data can be found in Table 2.

10.1371/journal.pone.0310830.g003 Fig 3 Comparison of anatomical and CT methods for measuring the eyeball and intraocular volume in New Zealand rabbits (ml).

Note: A, Comparison of two measurement methods in rabbit ocular volume; B, Comparison of two measurement methods in rabbit lens volume; C, Comparison of two measurement methods on the volume of rabbit anterior chamber; D, Comparison of two measurement methods on the volume of rabbit vitreous cavity (The volume of the vitreous cavity here represents the total volume of the vitreous cavity and the remaining eye tissue). ns means that the difference is not statistically significant; *Indicates P ≤ 0.05; *** Indicates P ≤ 0.001.

10.1371/journal.pone.0310830.t002 Table 2 Comparison of anatomical and CT measurements of ocular volumes and intraocular contents volumes in rabbits (ml).

Group (n = 12)	Ocular volume	Lens volume	Anterior chamber volume	Vitreous cavity volume (volume of vitreous cavity and remaining eye tissue)	Real vitreous cavity volume	
Gross volumetric estimation of ocular volume	3.04±0.24	0.44±0.02	0.26±0.05	2.34±0.26	1.54±0.14	
CT result	3.02±0.24	0.41±0.02	0.23±0.04	2.39±0.22	/	
t value	0.20	4.28	2.08	0.55	/	
P value	0.84	≤ 0.00***	0.05*	0.59	/	
Note:

*Indicates P ≤ 0.05;

*** Indicates P ≤ 0.001.

Results of ocular volume and intraocular contents in SD rats

There was no statistically significant difference (p>0.05) between the anatomical and CT measurements of eyeball volume and anterior chamber volume in SD rats, as demonstrated in Fig 4A and 4C. However, the CT measurements of lens volume were significantly larger than the gross volumetric estimation of ocular volume, as depicted in Fig 4B. Conversely, the gross volumetric estimation of ocular volume showed larger results for vitreous volume, as illustrated in Fig 4D. The eye volume, lens volume, anterior chamber volume and vitreous volume of SD rats are respectively about: 0.14±0.02 to 0.15±0.01ml, 0.03±0.00 to 0.03±0.00ml, 0.01±0.00 to 0.01±0.01ml, 0.04±0.01ml. Specific results for both measurement methods are detailed in Table 3.

10.1371/journal.pone.0310830.g004 Fig 4 Comparison of anatomical and CT methods for measuring the eyeball and intraocular volume in SD rats (ml).

Note: A, Comparison of two measurement methods in rat ocular volume; B, Comparison of two measurement methods in rat lens volume; C, Comparison of two measurement methods on the volume of rat anterior chamber; D, Comparison of two measurement methods on the volume of rat vitreous cavity (The volume of the vitreous cavity here represents the total volume of the vitreous cavity and the remaining eye tissue). ns means that the difference is not statistically significant; *Indicates P ≤ 0.05; ** Indicates P ≤ 0.01.

10.1371/journal.pone.0310830.t003 Table 3 Comparison of anatomical and CT measurements of ocular volumes and intraocular contents volumes in SD rats (ml).

Group (n = 12)	Ocular volume	Lens volume	Anterior chamber volume	Vitreous cavity volume (volume of vitreous cavity and remaining eye tissue)	Real vitreous cavity volume	
Gross volumetric estimation of ocular volume	0.15±0.01	0.03±0.00	0.01±0.01	0.11±0.01	0.04±0.01	
CT result	0.14±0.02	0.03±0.00	0.01±0.00	0.09±0.02	/	
t value	1.84	3.73	0.27	2.50	/	
P value	0.08	0.00**	0.79	0.02*	/	
Note:

*Indicates P ≤ 0.05;

** Indicates P ≤ 0.01.

Correlation analysis

The gross volumetric estimation of ocular volume results showed a significant positive correlation between the vitreous cavity volume and ocular volume in Bama miniature pigs (p ≤ 0.001, Fig 5A); The vitreous cavity volume of New Zealand rabbits was significantly positively correlated with the ocular volume (p ≤ 0.001), and significantly negatively correlated with the anterior chamber volume (p ≤ 0.05). The real vitreous volume was significantly positively correlated with the lens volume (p ≤ 0.05), as shown in Fig 5B; The vitreous cavity volume of SD rats was significantly positively correlated with the ocular volume (p ≤ 0.001), while the true vitreous volume was significantly negatively correlated with the ocular volume (p ≤ 0.05), as shown in Fig 5C.

10.1371/journal.pone.0310830.g005 Fig 5 Correlation heatmap of gross volumetric estimation of ocular volume data.

Note: A. Correlation analysis of Bama miniature pig data; Correlation analysis of New Zealand rabbit data; C. Correlation analysis of SD rat data. Red represents positive correlation, blue represents negative correlation.*P ≤ 0.05; **P ≤ 0.01; ***P ≤ 0.001.

As shown in Fig 6A, there was a significant positive correlation (p ≤ 0.001) between the vitreous cavity volume and ocular volume in the CT measurement results of Bama miniature pigs; The eyeball volume of New Zealand rabbits was significantly positively correlated with the vitreous cavity volume (p ≤ 0.001), and also significantly positively correlated with the lens volume (p ≤ 0.05), as shown in Fig 6B; The correlation of CT data in SD rats wasthe same as that in New Zealand rabbits, as shown in Fig 6C.

10.1371/journal.pone.0310830.g006 Fig 6 Correlation heatmap of Correlation heatmap of CT measurement data.

Note: A. Correlation analysis of Bama miniature pig data; Correlation analysis of New Zealand rabbit data; C. Correlation analysis of SD rat data. Red represents positive correlation, blue represents negative correlation.*P ≤ 0.05; **P ≤ 0.01; ***P ≤ 0.001.

Discussion

In this study, we employed micro-CT and anatomical methods to investigate the ocular volume and intraocular contents of Bama miniature pigs, New Zealand rabbits, and SD rats. Our findings contribute to the understanding of the measurement accuracy and potential applications of these techniques in ophthalmic research. Micro-CT, characterized by high resolution comparable to an optical microscope, provides a non-destructive means of studying internal microstructures [20], allowing for detailed anatomical studies of eye soft tissues [21]. Meanwhile, micro-CT can be applied in animal experiments to live small animals such as rats and rabbits, as well as to study various ex vivo tissue samples. In our study, due to the large size of the pigs, in vivo, eye scanning could not be performed. In order to control the uniform conditions, ex vivo CT scanning was performed on the eyeballs of pigs, rabbits, and rats. Our results revealed no statistically significant difference in ocular volume between CT and gross volumetric estimation of ocular volume across the three animal species. However, deviations in anterior chamber and lens volumes were observed. These discrepancies may arise from challenges in manually delineating the eyeball contour on CT images, potential tissue overlap, and subtle errors in gross volumetric estimation of ocular volume.

Based on the measurement results of the two methods, we observed similarities in pigs and rabbits, indicating that both species exhibited higher anterior chamber and lens volumes measured by anatomical methods compared to micro-CT, with statistically significant differences. Conversely, in SD rats, the anterior chamber volume measured by anatomical methods was smaller than that measured by micro-CT, while the vitreous volume was larger than that measured by CT. Despite the expectation of no difference in the volume of each eyeball content, this was not the case in reality. Compared to SD rats, the eyeballs of Bama miniature pigs and New Zealand rabbits are significantly larger, especially the eyeball volume of pigs, while the eyeballs of New Zealand rabbits are slightly smaller than those of pigs. We identified the main sources of error in obtaining eye content volume in pigs and rabbits using both measurement methods as the anterior chamber volume and lens volume. Micro-CT primarily utilizes scanning and manual delineation to obtain these volumes, reconstructing the anterior chamber and lens comprehensively and calculating their volumes. In contrast, gross volumetric estimation of ocular volume determine the volume of the anterior chamber through a single extraction of aqueous humor and a syringe scale. Therefore, we posit that CT measurement results may be more reliable because only the anterior chamber is delineated in the measurement, whereas the larger volume of the pig and rabbit anterior chamber measured by dissection may be due to the extraction of a small amount of posterior chamber volume.

We euthanized all animals by bleeding the abdominal aorta after anesthesia, and it took less than 2 minutes to completely kill the animals by cutting the abdominal aorta. Our procedure was to quickly transfer the extracted eyeballs into the CT room for scanning, with each eyeball scan lasting approximately 3–5 minutes. The scanned eyeball was first placed into the anatomical measurement volume for measurement and recording of the eyeball volume, which takes about 1–2 minutes. Then, the aqueous humor was immediately collected, which takes about tens of seconds. The eyeball that has been completely drained of aqueous humor was quickly frozen in liquid nitrogen, and the lens and vitreous body were removed and placed in the anatomical measurement container. After the tissue was thawed, readings were taken. We tried to extract the anterior chamber as dry as possible during the collection of aqueous humor, and evaluate all obtained parameters based on anatomical measurements and CT scans. We mentioned that the entire scanning process was very brief and there was almost no change in eye size, which is evidenced by the lack of statistical differences in the total eye volume of the three animals in CT and anatomical measurements. Regarding the smaller anterior chamber volume measured by dissection in SD rats compared to CT, SD rats have notably small eyeballs, necessitating higher accuracy in gross volumetric estimation of ocular volume. Additionally, their aqueous humor content is minimal, which may lead to losses during transfer to the measuring instrument. This discrepancy may be the primary reason why the anterior chamber volume measured by dissection is smaller than that measured by CT. The larger vitreous volume is likely due to this volume being mainly the sum of eyeball volumes minus the anterior and lens volumes. A smaller anterior chamber volume results in an overestimation of this portion of the volume. The loss error caused by the larger eyeball structure and contents of rabbits and pigs can be ignored, resulting in no difference between the two methods in vitreous volume.

Furthermore, the lens volumes of pigs and rabbits measured by dissection were both larger than the results of micro-CT. We do not attribute the main source of error to CT, as there is no discrepancy between the eye volume results obtained from CT and gross volumetric estimation of ocular volume without separating the intraocular contents. This suggests that we encounter no significant issues in delineating the eye structure, conducting 3D reconstruction, and automatically measuring volume. We speculate that the main source of error lies in gross volumetric estimation of ocular volume.

The process of anatomical measurement involves completely removing the lens after rapid freezing in liquid nitrogen and then placing it in a container for volume reading after thawing. This process includes freezing, separation, thawing, and data reading, each step potentially introducing errors. Particularly during the separation process, there may be some surrounding tissue included, leading to slightly larger measurement results. However, each lens we obtain remains frozen and intact until completely melted before reading. Additionally, the lens of SD rats is very small, and even if a small amount of surrounding tissue is included, it can be disregarded, resulting in no difference in lens volume between the two measurement methods. We maintain the view that gross volumetric estimation of ocular volume are essential because micro-CT may not clearly distinguish the boundary between the vitreous body and posterior chamber. We rely on gross volumetric estimation of ocular volume to accurately assess the volume of the vitreous body. In reality, we are not comparing two distinct measurement methods, but rather integrating these approaches to derive the volume of the eyeballs and their contents in Bama miniature pigs, New Zealand rabbits, and SD rats. We consider the results obtained from both methods as complementary experimental techniques rather than establishing comparative relationships between them. Therefore, we believe that referencing the results obtained from both methods together is necessary for a comprehensive understanding.

Due to limitations in experimental conditions, the small animal imaging technology available in our laboratory is micro-CT. In fact, possessing micro-CT equipment is considered rare and luxurious for many laboratories. Among the imaging techniques used in ophthalmology, ultrasound imaging (US), CT, and MRI are commonly used. The US uses ultrasound for disease diagnosis, which has the advantages of high soft tissue resolution [22], non-invasive, real-time imaging [23], and low cost [24]. However, its examination accuracy is insufficient, small lesions are easily overlooked, and it does not have a multiplanar imaging function, making it impossible to estimate volume. Besides, MRI has better soft tissue imaging capabilities, enabling 3D stereoscopic imaging, which is non-destructive and radiation-free for patients [25]. Compared with conventional US, MRI can more accurately measure small tumors [26]. However, due to the influence of metals, it cannot be used to measure tissues containing metals, and the operation time is long, with high usage costs and expenses [27, 28]. Compared to the two-dimensional imaging of the US, both CT and MRI, being three-dimensional imaging techniques, are suitable for volume estimation and are unaffected by metallic elements [29]. The examination time is short, the cost is low, one study found that using abdominal CT examination instead of direct MRI examination for lumbar diseases can save $1.2–3.4 billion per year [30]. And it is also a non-invasive operation [31], although CT radiation is relatively high, it is not significantly affected in animal experiments.

Micro-CT has the same imaging principle as clinical CT [32], but its resolution is higher [33], which is widely used for obtaining animal imaging data [34]. The imaging principle of Micro-CT mainly uses X-rays [35] to pass through the sample and capture hundreds of two-dimensional images from different angles around the sample. The software can reconstruct images from various angles to generate three-dimensional images [36], and it can also outline the parts of interest in the sample, and even create 3D animations. In addition, micro-CT can measure the size, volume, and spatial coordinates of each point of the sample, as well as the attenuation value and density information of the sample. In the study, we used micro-CT to scan the eyeballs of Bama miniature pigs, New Zealand rabbits, and SD rats. We manually delineated the anterior chamber and lens using software to construct complete stereoscopic images of the eyeballs, and successfully obtained the volumes of various regions of interest. However, the gap between the vitreous cavity and the posterior chamber on CT images is unclear, making it difficult to manually sketch the vitreous cavity, which is also one of the reasons why we combined gross volumetric estimation of ocular volume.

In fact, in order to prevent the unclear structure of the eyeball during the experimental design, we wanted to enhance image contrast by combining staining and micro-CT scanning. Like Leszczyński B, et al. [14] stained the eyeballs of domestic pigs with iodine, 100% Lugol, phosphotungstic acid, and 1% osmium tetroxide solutions, and then performed micro-CT scans. They found that staining can improve the visualization of eyeball structure, and different staining agents and staining time can affect the imaging results. However, the eyeball of SD rats is very small, and we are afraid that injecting staining agents may increase the intraocular volume. Therefore, we decided to dissect the eyeball after CT scanning separately to obtain its anatomical data, and estimate the volume of the eyeball and its contents by combining CT and gross volumetric estimation of ocular volume. Notably, the combination of micro-CT and gross volumetric estimation of ocular volume in this study allowed for comprehensive data acquisition, although challenges were encountered in delineating the vitreous cavity due to unclear boundaries. Future improvements could involve enhancing image contrast through staining. Our study lays the foundation for estimating eye tissue volumes in Bama miniature pigs, New Zealand rabbits, and SD rats, offering insights for drug research and intraocular drug volume conversion in these experimental animals.

Our correlation analysis results indicated a significant positive correlation (p ≤ 0.001) between the ocular volume and the vitreous cavity volume, regardless of anatomical or CT measurements, this may be mainly because the volume of the vitreous cavity was the sum of the volume of the real vitreous cavity and the remaining eye tissue, which was closer to the volume of the eyeball. But in the results of gross volumetric estimation of ocular volume. For example, we found that the vitreous cavity volume of New Zealand rabbits had a significant negative correlation with the anterior chamber volume, while the real vitreous cavity volume had a significant positive correlation with the lens volume. There was a significant negative correlation between the true vitreous cavity volume and eyeball volume in SD rats, which did not exist in the data of pig eyeball. We speculated that this might be due to the difference in the proportion of different eye contents in pigs, rabbits and SD rats (The lens of SD rats accounts for 18% of the total volume of the eyeball, while the true vitreous cavity accounts for 28%; the crystalline lens of rabbits accounts for 15% of the total volume of the eyeball, while the true vitreous cavity accounts for 50%; and the crystalline lens of pigs accounts for 6% of the total volume of the eyeball, while its true vitreous cavity accounts for 58%.). However, due to imaging defects in micro-CT, CT measurements cannot obtain the true volume of the vitreous cavity, so the correlation analysis results between the three different animal eye parameters measured by CT were almost identical.

So far, no other studies have reported on the volume of eyeballs and their contents in various structures of Bama miniature pigs, New Zealand rabbits, or SD rats using combined anatomical and imaging measurement methods, as observed in our study. However, some studies have utilized micro-CT to assess the volume of intraocular structures in animals. For instance, Atwood RC et al. [37] employed three-dimensional reconstruction via micro-CT and computer-aided image analysis technology to quantify the volume of intraocular vessels in mice. Similarly, Leszczynski et al. [14] successfully utilized micro-CT to obtain data on pig eyeball volume. Diverging from the aforementioned studies, which focused on quantifying the volume of eye contents at specific sites, our study aimed to elucidate the volume of eyeballs and their contents across several commonly used experimental animals. This approach aimed to provide practical reference values for the utilization of relevant experimental drugs and the conversion of drug volume between species. Therefore, we conducted comprehensive measurements of the volume of the anterior chamber, lens, and vitreous body. Nevertheless, our research is built upon the foundation laid by previous studies, which affirmed and elucidated the value of micro-CT and its 3D reconstruction technology in measuring eyeball volume. These earlier investigations paved the way for our study’s methodology and its contributions to the field.

Despite its contributions, our study has several limitations. Firstly, measurement errors, particularly inherent to the traditional anatomical measurement methods we employed, remain a challenge. Despite our efforts to enhance the accuracy of each measurement step, errors, especially in measuring anterior chamber and lens volume, are inevitable. We anticipate that future advancements in measurement technology may mitigate these issues. Furthermore, the indistinct boundary between the vitreous cavity and posterior chamber in micro-CT images presents a significant challenge. While we attempted to optimize imaging parameters to achieve clearer images, none proved successful. Additionally, we hesitated to use staining techniques due to concerns about increasing intraocular volume. Access to equipment with clearer resolution, such as micro-MRI or high-resolution micro-CT, could potentially offer solutions. However, our current laboratory lacks the resources to acquire such equipment.

Conclusion

Our study demonstrates the feasibility of estimating the volume of the eyeball and its contents through a combined approach utilizing micro-CT and anatomical methods in Bama miniature pigs, New Zealand rabbits, and SD rats. Notably, we observed variations in the proportion of eye contents relative to the eyeball volume across different species. Specifically, SD rats exhibited the largest proportion of lens volume in the total eyeball volume, emphasizing the potential vulnerability of the lens during intraocular procedures in this species. Further refinements and technological advancements are crucial for enhancing precision and expanding the applications of these techniques in ophthalmic research.

Supporting information

S1 Data (ZIP)

S2 Data (ZIP)

S3 Data (ZIP)

S4 Data (ZIP)

Thanks to WestChina-Frontier PharmaTech Co., Ltd. for providing laboratory facilities and animals for this study.

10.1371/journal.pone.0310830.r001
Decision Letter 0
Goździewska-Harłajczuk Karolina Academic Editor
© 2024 Karolina Goździewska-Harłajczuk
2024
Karolina Goździewska-Harłajczuk
https://creativecommons.org/licenses/by/4.0/ This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Submission Version0
27 Aug 2024

PONE-D-24-29172Anatomical and Micro-CT measurement analysis of ocular volume and intraocular volume in adult Bama Miniature Pigs, New Zealand rabbits, and Sprague-Dawley ratsPLOS ONE

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Reviewer #2: Yes

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Reviewer #1: The study aimed to measure the eyeball and its components in Bama Miniature Pigs, New Zealand rabbits, and Sprague-Dawley rats using micro-CT and anatomical techniques. They included six animals from each species and measured the volumes of the eyeball, anterior chamber, lens, and vitreous cavity. The conclusion was that combining micro-CT and anatomical measurements is effective for determining eyeball volumes in these animals. This information could be useful for future ophthalmology-related drug research.

Reviewer #2: Review on the manuscript PONE-D-24-29172

Anatomical and Micro-CT measurement analysis of ocular volume and intraocular

volume in adult Bama Miniature Pigs, New Zealand rabbits, and Sprague-Dawley rats

Thanks very much for giving me such opportunity to revise the current manuscript. The manuscript describes the anatomical and micro-CT examination of the eye in Bama miniature pigs, New Zealand rabbits and Sprague-Dawley rats.

The manuscript could be considered for publication. My main concern is that the study design is not clear, was it an in vivo or ex vivo or in vitro (as indicated by authors). It is not clear when animals really euthanized (before or after CT scan). If before scanning, how did the author maintain accuracy of measurements following euthanasia?

Additionally, the method of euthanasia requires justification.

Minor comments:

Title:

- Replace “Pig” with “pig”.

Abstract:

- You can report your results using 2 decimal points.

Introduction:

- Line 78: “Salguero R et al.”…replace with “Salguero et al.”

- “length of each line in normal dog”….explain what do you mean by each line.

Method:

- Lone 89: “This is an in vitro experiment”…..Do you believe that this is an in vitro experiment?! ….. please replace by “ an in vivo study”.

- Line 103: “The tool was then used to draw the ROI (region of interest)” could be replaced by “The tool was then used to draw the region of interest (ROI)”

- Line 110: “All operations were performed by a single experimenter” … Do you mean “All CT scans and measurements were made by the same examiner (add Name abbreviations)?? ….Did you made measurements at the same time of examination or you made it on saved scans….please clarify. Did you make your measurements only one time or multiple times and get your mean for each eye measurements.

- Line 111: Anatomical results/anatomic measurements all over the text could be replaced by “Gross volumetric estimation of ocular volume”

- Line 112-113:

- “intravenous injection of pentobarbital sodium (30mg/kg)” at which vein did you inject in each animal?

- please add concentration, trade name, manufacturer and country of origin of the anesthetic drugs.

- “all animals were euthanized through abdominal artery bleeding” ….Which abdominal artery? How did you approach it? What is your justification and reference for this method of euthanasia? How did you ensure euthanasia?

Statistical analysis:

- Did you consider running correlation and regression analysis to your measurements?

Results:

- Results could be presented using 2 decimal points.

Discussion:

- Line 162-163: “In our study, due to the large size of the pigs, in vivo, eye scanning could not be performed. In order to control the uniform conditions, ex vivo CT scanning was performed on the eyeballs of pigs, rabbits, and rats”…..this is really very confusing. For the first time here in the discussion section to include that you did not make your CT scans on life animals??!!! You have mentioned that animals were euthanized for the anatomic study not for the CT scans.

Then how did you position the eyes balls for scanning? How did you manage post-mortem changes of ocular dimensions during scanning??

Please clarify in details.

Line 173: you are suggesting that the micro-CT measurements are more reliable than anatomic measurements “we posit that CT measurement results may be more reliable”……So what was your study objective and hypothesis??!! What is your gold standard to judge/prove that micro CT is reliable in determination of volumetric eye measurement?

Figures:

- Figure 1: please add reference to panels a,b, c within “Figure caption”. You may need to include description of volumetric measurements in the CT scan.

Thanks

**********

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Reviewer #1: Yes: Rodrigo Pessoa Cavalcanti Lira

Reviewer #2: No

**********

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10.1371/journal.pone.0310830.r002
Author response to Decision Letter 0
Submission Version1
31 Aug 2024

Dear Editor and reviewers, thank you very much for giving us the opportunity to revise this manuscript. We have carefully revised the manuscript according to your requirements, and the relevant response is as follows. Once again, we sincerely thank you!

To academic editor and journal:

1.Please ensure that your manuscript meets PLOS ONE's style requirements, including those for file naming.

Reply:

Dear editor, thank you again for providing us with this opportunity to revise the manuscript. We have changed the revised manuscript into the correct format as required by the journal, please check it!

2. Thank you for stating the following financial disclosure:

“2023 Research Fund of Aier Eye Research Institute(No.AEI202310LC01); Science Research Foundation of Aier Eye Hospital Group (No.AR2201D3).” Please state what role the funders took in the study. If the funders had no role, please state: "The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript." If this statement is not correct you must amend it as needed. Please include this amended Role of Funder statement in your cover letter; we will change the online submission form on your behalf.

Reply:

Dear editor, thank you very much for your comment, we are very sorry for not stating the role of funders in the cover letter, we have added relevant information in the cover letter, thank you very much for your comments again!

3.Thank you for stating the following in the Acknowledgments Section of your manuscript:

“Thanks to Aier Eye Group for its funding support; Thanks to WestChina-Frontier PharmaTech Co., Ltd. for providing laboratory facilities and animals for this study.”We note that you have provided funding information that is currently declared in your Funding Statement. However, funding information should not appear in the Acknowledgments section or other areas of your manuscript. We will only publish funding information present in the Funding Statement section of the online submission form.

Reply:

Dear editor, thank you very much for your comment. We have deleted the funding information from the manuscript and added it in the cover letter, please check it!

4.In the online submission form you indicate that your data is not available for proprietary reasons and have provided a contact point for accessing this data. Please note that your current contact point is a co-author on this manuscript. According to our Data Policy, the contact point must not be an author on the manuscript and must be an institutional contact, ideally not an individual. Please revise your data statement to a non-author institutional point of contact, such as a data access or ethics committee, and send this to us via return email. Please also include contact information for the third party organization, and please include the full citation of where the data can be found.

Reply:

Dear editor, thank you very much for your comment. We wanted to submit the original data when submitting the manuscript, but the original data was too large to be submitted. That's why we wrote in the data statement that we can obtain the original data from the corresponding author. In this revised draft, we have split the original data and uploaded it, please check, thank you!

5.Please review your reference list to ensure that it is complete and correct. If you have cited papers that have been retracted, please include the rationale for doing so in the manuscript text, or remove these references and replace them with relevant current references. Any changes to the reference list should be mentioned in the rebuttal letter that accompanies your revised manuscript. If you need to cite a retracted article, indicate the article’s retracted status in the References list and also include a citation and full reference for the retraction notice.

Reply:

Dear editor, thank you very much for your comment. The references cited in this manuscript are complete and correct, and there are no references to the retracted manuscript.

To additional editor:

Based on the Reviewers` comment the current manuscript needs some minor revision.

Reply:

Dear editor, thank you again for providing us with this opportunity to revise the manuscript. We have made corresponding revisions to the manuscript as required by the reviewers, please check it! Thanks again for this opportunity！

To reviewers:

To Reviewer #1:

The study aimed to measure the eyeball and its components in Bama Miniature Pigs, New Zealand rabbits, and Sprague-Dawley rats using micro-CT and anatomical techniques. They included six animals from each species and measured the volumes of the eyeball, anterior chamber, lens, and vitreous cavity. The conclusion was that combining micro-CT and anatomical measurements is effective for determining eyeball volumes in these animals. This information could be useful for future ophthalmology-related drug research.

Reply:

Dear reviewer, thank you very much for your detailed review of this manuscript. Your comments are an important recognition and support for our work. Thank you again for your comments.

To Reviewer #2:

1.Title:

- Replace “Pig” with “pig”.

Abstract:

- You can report your results using 2 decimal points.

Introduction:

- Line 78: “Salguero R et al.”…replace with “Salguero et al.”

- “length of each line in normal dog”….explain what do you mean by each line.

Reply:

Dear reviewer, thank you very much for your comments. We have made corresponding modifications in the manuscript. Besides, we are very sorry for the confusion caused by our writing, the "length of each line in normal dog" refers to the distance between certain structures of the dog's eyeball, including the average axial length of the sphere, the average anterior posterior distance of the anterior chamber, the average anterior posterior distance of the vitreous chamber, etc. For better understanding, we have replaced this sentence with “and the length of the normal dog eye structures (Including the average axial length of the sphere, the average anterior posterior distance of the anterior chamber, the average anterior posterior distance of the vitreous chamber, etc) with CT”, please check it.

2.Method:

- Lone 89: “This is an in vitro experiment”…..Do you believe that this is an in vitro experiment?! ….. please replace by “ an in vivo study”.

- Line 103: “The tool was then used to draw the ROI (region of interest)” could be replaced by “The tool was then used to draw the region of interest (ROI)”

Reply:

Dear reviewer, thank you very much for your comments. We have made corresponding modifications in the manuscript. Besides, due to our inability to perform micro CT scans on live pigs and rabbits, the scanning chamber for small animal CT is very small and can only accommodate mice and rats. In order to standardize the measurement method, we removed the eyeball and conducted separate CT and anatomical measurements. We euthanize animals in order to extract their eyeballs for measurement of in vitro parameters. Strictly speaking, we should not consider it as an in vivo experiment. To avoid misunderstandings, we have removed the mention of in vivo and in vitro experiments. We have provided corresponding descriptions of the measurement methods in our methodology.please check it.

3.Line 110: “All operations were performed by a single experimenter” … Do you mean “All CT scans and measurements were made by the same examiner (add Name abbreviations)?? ….Did you made measurements at the same time of examination or you made it on saved scans….please clarify. Did you make your measurements only one time or multiple times and get your mean for each eye measurements.

Reply:

Dear reviewer, thank you very much for your comment. In order to reduce errors, our CT scans and eye measurements under CT were performed by the same professional imaging technician (Zongtao Shu). Due to the time-consuming process of manual delineation and reconstruction of eye structure involved in the measurement process, our technicians promptly saved all eye image data after scanning, and then uniformly measured eye volume. In addition, we also referred to other literature and did not have similar imaging data for multiple measurements. Therefore, the CT measurement data for this part was only reconstructed by outlining the eyeball structure once to obtain the required parameter results. The entire process took nearly 2 months and was quite cumbersome and complex.

4.Line 111: Anatomical results/anatomic measurements all over the text could be replaced by “Gross volumetric estimation of ocular volume”.

Reply:

Dear reviewer, thank you very much for your comments. We have made corresponding modifications in the manuscript, please check it.

5.- Line 112-113:

- “intravenous injection of pentobarbital sodium (30mg/kg)” at which vein did you inject in each animal?

- please add concentration, trade name, manufacturer and country of origin of the anesthetic drugs.

Reply:

Dear reviewer, thank you for your detailed comments on this manuscript. Both Bama miniature pigs and New Zealand rabbits were injected via ear vein after skin preparation, while SD rats were injected intraperitoneally with 30mg/kg pentobarbital sodium. We are very sorry that we did not provide detailed information on the concentration, product name, manufacturer, and country of origin of the anesthetic drug in the manuscript. Our reagents are 2mg/kg diazepam (Sigma-Aldrich, USA) and 30mg/kg pentobarbital sodium (Sigma-Aldrich, USA), which we have supplemented in the manuscript. Please check. Thank you!

6.“all animals were euthanized through abdominal artery bleeding” ….Which abdominal artery? How did you approach it? What is your justification and reference for this method of euthanasia? How did you ensure euthanasia?

Reply:

Dear reviewer, thank you very much for your comments. All animals in this manuscript were euthanized with diazepam and anesthetized with 30mg/kg pentobarbital sodium before cutting the abdominal aorta and bleeding to death. We have supplemented this information in the manuscript. We ensure that all animals have a safe, effective, painless, and pain free death experience, following the AVMA Animal Euthanasia Guidelines. In addition, we confirmed whether the signs of life had completely disappeared by observing the disappearance of the animal's breathing and heartbeat, as well as pinching its toe reflex. After confirming that the animal had completely died, we proceeded to remove the eyeball, which we also supplemented in the manuscript. Finally, thank you again for your comments!

7.Statistical analysis:

- Did you consider running correlation and regression analysis to your measurements?

Reply:

Dear reviewer, thank you very much for your valuable comment. We have added relevant statistics to the manuscript, please check it!

8.Results:

- Results could be presented using 2 decimal points.

Reply:

Dear reviewer, thank you very much for your comments. We have made corresponding modifications in the manuscript, please check it.

9.Discussion:

- Line 162-163: “In our study, due to the large size of the pigs, in vivo, eye scanning could not be performed. In order to control the uniform conditions, ex vivo CT scanning was performed on the eyeballs of pigs, rabbits, and rats”…..this is really very confusing. For the first time here in the discussion section to include that you did not make your CT scans on life animals??!!! You have mentioned that animals were euthanized for the anatomic study not for the CT scans.

Then how did you position the eyes balls for scanning? How did you manage post-mortem changes of ocular dimensions during scanning??

Please clarify in details.

Reply:

Dear reviewer, thank you very much for your valuable comments. We are very sorry that our manuscript has caused you inconvenience. In fact, we have written about the specific process in the measurement of eye volume in methodology: we euthanize all animals by bleeding the abdominal aorta after anesthesia, and it takes less than 2 minutes to completely kill the animals after cutting the abdominal aorta. Our procedure is to quickly transfer the extracted eyeballs into the CT room for scanning, with each eyeball scan lasting approximately 3-5 minutes. The scanned eyeball is first placed into the anatomical measurement volume for measurement and recording of the eyeball volume, which takes about 1-2 minutes. Then, the aqueous humor is immediately collected, which takes about tens of seconds. The eyeball that has been completely drained of aqueous humor is quickly frozen in liquid nitrogen, and the lens and vitreous body are removed and placed in the anatomical measurement container. After the tissue is thawed, readings are taken. We try to extract the anterior chamber as dry as possible during the collection of aqueous humor, and evaluate all obtained parameters based on anatomical measurements and CT scans.

We are unable to perform micro-CT scans on live pigs and rabbits because the scanning chamber for small animal-CT is very small and can only accommodate mice and rats. In order to standardize the measurement method, we removed the eyeball and conducted separate CT and anatomical measurements. We euthanize animals in order to extract their eyeballs for measurement of in vitro parameters. Strictly speaking, we should not consider it as an in vivo experiment. To avoid misunderstandings, we have removed the mention of in vivo and in vitro experiments. We have provided corresponding descriptions of the measurement methods in our methodology. In addition, we mentioned that the entire scanning process was very brief and there was almost no change in eye size, which is evidenced by the lack of statistical differences in the total eye volume of the three animals in CT and anatomical measurements.

Sorry, we did not provide a detailed explanation of the specific method for locating the eyeball for scanning. We have added it to the methodology (The specific way we locate the eyeball for scanning is to ensure that the entire eyeball is located within the scanning interval, manually select the lens to start drawing from the tip of the ciliary body, move towards the other end until the lens structure disappears, and then use this tool to draw the region of interest (ROI) of the lens until the anatomical structure disappears; Using the same method, select any starting surface of the eyeball or anterior chamber structure and draw its ROI until the anatomical structure disappears,the vitreous body and posterior chamber cannot be located due to the lack of a clear boundary line), please check it. Finally, thank you again for your valuable comments. Thank you!

10.Line 173: you are suggesting that the micro-CT measurements are more reliable than anatomic measurements “we posit that CT measurement results may be more reliable”……So what was your study objective and hypothesis??!! What is your gold standard to judge/prove that micro CT is reliable in determination of volumetric eye measurement?

Reply:

Dear reviewer, thank you very much for your detailed review of this manuscript. We are pleased with your professional feedback. We are very sorry that we have not actually recognized micro CT as the gold standard for measuring eye volume. We have also mentioned in our discussion that MRI is actually a more accurate measurement method, but small animal MRI is a very luxurious existence in our laboratory, and currently we do not have the conditions to implement it. The main reason why we believe CT results are more reliable is as follows: we found that there were differences between the anterior chamber volume of the eyeballs of three animals and the CT results. For example, the anatomical measurement of the anterior chamber volume in pigs was 0.28±0.04ml, while the CT result was 0.19±0.05ml. We can only combine the results of the two methods to conclude that the anterior chamber volume in pigs is approximately 0.19±0.05-0.28±0.04ml. We are not sure if time will have a certain impact on the amount of room water, but there are indeed differences between these two measurement methods on room water. In fact, we believe that CT measurement results are more reliable becaus

Attachment Submitted filename: Response to Reviewers.docx

10.1371/journal.pone.0310830.r003
Decision Letter 1
Goździewska-Harłajczuk Karolina Academic Editor
© 2024 Karolina Goździewska-Harłajczuk
2024
Karolina Goździewska-Harłajczuk
https://creativecommons.org/licenses/by/4.0/ This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Submission Version1
8 Sep 2024

Anatomical and Micro-CT measurement analysis of ocular volume and intraocular volume in adult Bama Miniature pigs, New Zealand rabbits, and Sprague-Dawley rats

PONE-D-24-29172R1

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10.1371/journal.pone.0310830.r004
Acceptance letter
Goździewska-Harłajczuk Karolina Academic Editor
© 2024 Karolina Goździewska-Harłajczuk
2024
Karolina Goździewska-Harłajczuk
https://creativecommons.org/licenses/by/4.0/ This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
11 Sep 2024

PONE-D-24-29172R1

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